Compositions and Methods for the Treatment of Diseases Associated with Aberrant Cilia Assembly and Regulation

ABSTRACT

Compositions and methods are provided for identifying agents which have efficacy for the treatment of disorders related to aberrant cilial structure and function, including polycystic kidney disease.

This application is a Continuation-in-Part Application of U.S. patent application Ser. No. 12/374,209 filed Apr. 2, 2009, which is a §371 Application of PCT/US07/73722, filed Jul. 17, 2007, which claims priority to U.S. Provisional Applications 60/831,479 and 60/925,272 filed Jul. 17, 2006 and Apr. 19, 2007. This CIP application also claims priority to U.S. Provisional Application, 61/122,303 filed Dec. 12, 2008. The entire disclosures of each of the foregoing application are incorporated herein by reference.

Pursuant to 35 U.S.C. §202(c), it is acknowledged that the U.S. Government has certain rights in the invention described, which was made in part with funds from the National Institutes of Health, Grant Numbers RO1 CA63366, CA-06927, DOD W81XWH-07-1-0676 and RO1 CA-113342.

FIELD OF THE INVENTION

The present invention relates to the fields of molecular biology and cilia-associated structural and cellular signal transduction. More specifically, the invention provides methods for identifying compounds which modulate cilia assembly and disassembly, thereby providing treatment for disorders associated with aberrant coordination of cilia function, including, for example, polycystic kidney disease, renal cysts, cancer, hypertension and infertility.

BACKGROUND OF THE INVENTION

Several publications and patent documents are cited throughout the specification in order to describe the state of the art to which this invention pertains. Each of these citations is incorporated by reference herein as though set forth in full.

In polycystic kidney disease (PKD), Bardet-Biedl Syndrome (BBS), and other disorders, mutations in cilia-associated structural or signaling proteins cause insensitivity to external mechanical and diffusible signaling cues, resulting in disorganized, hyperplastic cell growth (Benzing and Walz, 2006; Pan et al., 2005; Singla and Reiter, 2006). On the organismal level, ciliary defects produce renal cysts, infertility, respiratory disorders, situs inversus, and predisposition to obesity, diabetes, and hypertension. Notably, recent studies have shown that the Hedgehog, Wnt, PDGFαα, and other signaling cascades are coordinated at cilia (Cano et al., 2004; Huangfu and Anderson, 2005; Liu et al., 2005; Schneider et al., 2005; Simons et al., 2005; Tanaka et al., 2005). The frequent deregulation of these pathways during cell transformation, together with the common disappearance of cilia in transformed cells, raises the possibility that defective ciliary signaling may promote cancer.

Although an increasing number of proteins are being defined as ciliary structural components or cilia-associated signaling proteins, very little is currently known about the cellular machinery controlling the formation and resorption of cilia. It has long been known that cilia are regulated dynamically throughout the cell cycle. In many cells, resorption occurs at mitotic entry, and reappearance after progression into G1. However, resorption is not solely linked to mitotic entry, with some cells undergoing waves of resorption at different points in cell cycle: for example, Tucker et al. have noted ciliary resorption as cells emerge from quiescence, prior to S-phase (Quarmby and Parker, 2005; Rieder et al., 1979; Tucker et al., 1979). Given their increasingly apparent role in detecting and transmitting extracellular signals, regulated formation, disassembly, or shortening of cilia may play an important role in cellular growth controls, serving as a rheostat to limit response to overly persistent or abnormal cell growth cues in the extracellular environment.

A cilium arises from a basal body, a structure that differentiates from one of the centrioles in the centrosome in non-proliferating cells and organizes the microtubule bundles that constitute the ciliary axoneme. Cilia are evolutionarily related to the motile flagella of lower eukaryotes, such as the green algae Chlamydomonas. Genetic studies in Chlamydomonas have recently begun to dissect the process of flagellar resorption (Bradley and Quarmby, 2005; Marshall et al., 2005; Pan and Snell, 2005; Quarmby, 2004). These studies have identified altered functionality of the intraflagellar transport (IFT) machinery and destabilization of the axoneme as hallmarks of disassembly, and implicated CALK and other kinases as regulators of disassembly. The means by which CALK becomes activated at initiation of disassembly and the critical CALK effectors in the disassembly process remain unknown, as does the relevance of these observations to higher eukaryotes.

CALK is very distantly related to the human Aurora A (AurA) kinase, with 55% similarity centered on the protein catalytic domain. In humans, Aurora A (AurA) is a centrosomal kinase that regulates mitotic entry through activation of Cdk1-cyclin B and other substrates that organize the mitotic spindle (Bischoff et al., 1998; Marumoto et al., 2005). AurA amplification or activation is common in many cancers characterized by centrosomal amplification and genomic instability (Anand et al., 2003; Goepfert et al., 2002; Gritsko et al., 2003). In the past year, altered expression of the HEF1 (Law et al., 1996; O'Neill et al., 2000) scaffolding protein has recently been identified as part of a pro-metastatic signature in breast cancer (Minn et al., 2005), shown to contribute to the aggressiveness of glioblastomas (Natarajan et al., 2006), and found to be critical for progression to metastasis in melanomas (Kim et al., 2006). HEF1 is best known as a transducer of integrin-initiated attachment, migration, and anti-apoptotic signals at focal adhesions (O'Neill et al., 2000).

SUMMARY OF THE INVENTION

In accordance with the present invention, methods for identifying agents which modulate ciliary function and assembly/disassembly are provided. An exemplary method entails providing cells which express AurA and HEF1 and incubating the cells in the presence and absence of the agent. Following treatment, the cilia present on the cells are assessed for alterations which occur in the presence but not the absence of the agent, agents which cause alterations being identified as modulators of ciliary function and assembly.

In yet another aspect of the invention, an in vivo model for assessing agents which modulate kidney cyst formation is provided. In the model, a first strain of mice in which Pkd1 is conditionally inactivated is provided and the mice crossed with each of the following mice, i) a transgenic HEF-1 knock out mouse; ii) a mouse expressing functional HEF-1 and iii) a third strain of mice which are heterozygous for HEF-1 expression. After crossing, Pkd1 is inactivated and a test agent administered to each of the newly created strains of mice. After a suitable time period of administration, the mice are assessed to determine whether the agent modulates cyst formation relative to untreated mice.

In yet another aspect the invention provides a method for identifying agents which modulate calmodulin-Aurora A complex formation having efficacy for the treatment of polycystic kidney disease (PKD). An exemplary method entails providing kidney cells which express Aurora A and calmodulin and incubating cells in the presence and absence of said agent and in the presence of a molecule which induces release of Ca²⁺. The extent to which the agent disrupts calmodulin-Aurora A complex formation following release of calcium is then determined, agents which disrupt complex formation identified by the aforementioned method should have efficacy for the treatment of PKD. In a preferred embodiment, the agent inhibits activation of aurora A kinase and includes, without limitation, siRNA which specifically down modulate Aurora A expression, C1368, PHA-680632, MK-0457, ZM447439, MLN 8237, MLN8054, VX-680 and hesparadin. In further embodiments of the invention, the effect of the agent on disruption of aurora A binding to PC2 and/or cellular localization of PC2 and/or Aurora A phosphorylation of PC2 may be determined.

In yet another embodiment of the invention, a method for inhibiting progression of polycystic kidney disease in a patient in need thereof is provided. In one aspect the method entails administration of an effective amount of an aurora kinase inhibitor the patient in an amount effective to inhibit cyst formation in the kidney. The inhibitor may inhibit aurora kinase A or aurora kinase B or it may inhibit both.

BRIEF DESCRIPTION OF THE DRAWINGS

FIG. 1. Activation of AurA at the basal body occurs during ciliary disassembly. A. Assembly of cilia. An average of 200 cells were counted in two independent experiments B. Disassembly of cilia induced by serum stimulation. An average of 150 cells were counted in each of 4 experiments. C. Immunofluorescence of quiescent cells with antibody to AurA (green), acetylated α-tubulin (blue), and DNA (red). Scale bar 10 μm. In this and subsequent panels, boxes in main image indicate structures shown at high magnification to right. D. Immunofluorescence of quiescent cells with polyclonal rabbit antibody to HEF1 (green), also visualizing acetylated α-tubulin (blue), and DNA (red); compare also to E. Scale bar 10 μm. E. Immunofluorescence of quiescent cells with monoclonal antibody to HEF1 (green), also visualizing γ-tubulin (blue) and DNA (red). Scale bar 5 μm. See also FIG. 6A. F. Immunofluorescence of quiescent cells with antibody to phospho-AurA (green), acetylated α-tubulin (blue), and DNA (red). Scale bar 12.5 μm. G. Immunofluorescence of serum-stimulated cells with antibody to phospho-AurA (green), acetylated α-tubulin (blue), and DNA (red). Scale bar 5 μm. H. Western analysis of AurA and HEF1 in hTERT-RPE1 cells after serum stimulation. Western blots shown represent strips and reprobes of a single gel. Higher molecular weight HEF1 band reflects hyperphosphorylation, and coincides with AurA activation and ciliary disassembly at 2 and 24 hours after serum addition (at time point 0). Light gray arrow indicates cross-reactivity of phospho-AurA directed antibody with total AurA; black arrow indicates phospho-AurA. See also FIG. 2H. I. Immunofluorescence depicting AurA activation in serum-stimulated cells during disassembly of cilia. All images are merged panels of acetylated α-tubulin (red), phospho-AurA or total AurA (green) and DNA (blue).

FIG. 2. Profile of serum induced disassembly of cilia in hTERT-RPE1, IMCD3, and Caki-1 cells. A. Immunofluorescence analysis of cilia in quiescent hTERT-RPE1 cells. Cilia are visualized with antibodies to acetylated α-tubulin (Ac-α-tub, blue). The basal body and the second cellular centriole are visualized with antibodies to γ-tubulin (γ-tub, green), while DNA is indicated in red (PI). Insets are enlarged view of structures marked with arrowhead. Scale bar 10 μm. Note, some later analyses (FIGS. 8, 10) use glutamylated α-tubulin as a second marker for cilia. B. FACS profile of cells at hours (h) indicated after serum treatment of quiescent hTERT-RPE1 cells. Approximate 2N and 4N peaks are marked; cells at 24 hours show marked reduction of G1, and increased G2/M compartmentation. Cells >4N reflect aggregated cells, as well as a very small population that have failed cytokinesis. C. Quantification of BrDU-staining in cells confirms the peak of S-phase occurs at ˜42 hours after treatment of cells with serum. D., E. Time course of disassembly of cilia induced by serum stimulation in IMCD3 (D) and Caki-1 (E) cell lines. F., G. Western blot with antibodies to HEF1, AurA, and T288-phospho-AurA (phAurA) at 0, 2, and 18 hours after serum stimulation (see D, E) in IMCD-3 (F) and Caki-1 (G) cells. H. Experiment as in FIG. 1H, except using Cell Signaling antibody to visualize phospho-AurA. I. Experiment as in FIG. 3E, except using Cell Signaling antibody to visualize phospho-AurA

FIG. 3. Activation of AurA is necessary for ciliary resorption. A. Disassembly of cilia in cells treated with siRNA to AurA or HEF1, or with Scrambled (Scr) control siRNA, for 0 to 24 hours after serum addition. Assay performed 3 times, with an average of 100 cells counted/experiment by acetylated tubulin staining. Results were confirmed using a second antibody (anti-glutamylated tubulin) to independently score cilia following depletion (FIGS. 8D, 10E). B. Ciliary disassembly was induced in ciliated cells pre-treated with control (Scr), AurA-targeted (siA), or HEF1-targeted (siH) siRNA by supplementing growth media with serum. At 2, 12, and 24 hours after addition of serum, AurA was immunoprecipitated and used for an in vitro kinase assay as in (Pugacheva and Golemis, 2005). Shown, ³²P-labelled phosphorylated histone H3 (top) and total histone H3 in the reaction (stained with Coomassie Blue, bottom). C. Length of cilia in untreated hTERT-RPE1 cells (—), or the hTERT-RPE1 cells treated with control (Scr) or HEF1 targeting siRNA, at the indicated time points. D. Ciliated hTERT-RPE1 cells were treated with AurA inhibitor (PHA-680632) or DMSO, then disassembly of cilia tracked for 24 hours post serum addition. The in vitro IC50 of PHA-680632 is 27 nM for AurA; this compound also less potently inhibits AurC, AurB, and FGFR1 (IC50 120, 185, and 390 nM, respectively, (Soncini et al., 2006)). Results were confirmed using anti-glutamylated tubulin, as shown in FIG. 8D. E. Analysis performed in parallel with experiments described in D demonstrates PHA-680632 blocks appearance of T²⁸⁸-phospho-AurA (visualized with antibody from BioLegend), and HEF1 phosphorylation (115 kDa form), in reference to DMSO (−) at the 2 and 24 hour time points. Black arrows marks phosphorylated AurA, and hyperphosphorylated (p115) HEF1; gray arrow indicates p105 HEF1. See also FIG. 2I. F. Cells were treated with indicated concentrations of the AurA inhibitor PHA-680632, and then AurA immunoprecipitated, and used for in vitro kinase reactions (left) or whole cell lysates used for Western analysis with antibody to total or phosphorylated AurA (right). G. Immunofluorescence analysis of appearance of phospho-AurA at times indicated after serum stimulation in DMSO- or PHA-680632-treated cells. DNA (blue), acetylated α-tubulin (red), and T²⁸⁸-phospho-AurA (green). In 18 hr DMSO/ph-AurA, an asterisk (*) marks a rare observation of phospho-AurA at the base of a shortened cilium. H. FACS analysis of cells treated with DMSO vehicle or PHA-680632 at the times indicated after serum stimulation.

FIG. 4. Growth factor-induced disassembly of cilia in hTERT-RPE1 cells. A. Time course of disassembly of cilia in cells treated with EGF, PDGF, and TGF-β. To induce disassembly, hTERT-RPE1 cells were treated with EGF (10 ng/ml, Sigma), PDGF (10 ng/ml Sigma), and TGF-β (10 ng/ml, R&D Systems), and otherwise assayed as for serum treated cells. ***, one-way ANOVA indicates that reduction of cilia induced by PDGF is statistically significant (p<0.001) by 2 hours after treatment of cells. B. Western blot indicating T288-phosphorylated and total AurA in cells treated with PDGF or EGF, at times indicated after treatment. C. Immunofluorescence depicting cilia (red, acetylated a α-tubulin) and phospho-AurA (green) staining at the basal body at 2 hours after stimulation with PDGF, EGF, or serum. Phosphorylated AurA is apparent in PDGF- and serum-treated cells, but not EGF-treated cells: further, many well-formed cilia are apparent in EGF-treated cells (marked with asterisks) in contrast to other conditions. Scale bar represents 10 μM.

FIG. 5. Microinjection of active AurA causes rapid loss of cilia. A. Microinjection of wild type AurA, T288A or D274N mutant AurA, or GST, or PBS buffer, into hTERT-RPE1 cells with pre-formed cilia. (−), uninjected controls. Time reflects minutes from injection to initiating fixation of slides. Experiments repeated 3 times, with >100 injected cells scored in each experiment. B. Cilia 45 minutes post-injection of AurA or D274N. Red, acetylated α-tubulin; blue, glutamylated α-tubulin (a second independent marker of cilia); blue, DNA; green, Dextran488 indicates injected cells. High magnification images to right are from boxed cells; * marks magnification of uninjected cells. C. AurA and mutants (D274N, T288A) were incubated with histone H3 (17 kD) and MBP (22 kD) substrates in an in vitro kinase assay, confirming the activity of kinase. +Lysate indicates that mutants were incubated for 3 hours at 4° C. with hTERT-RPE1 cell lysate, then pulled down and used for the kinase assay.

FIG. 6. A. Besides the novel localization of HEF1 at cilia described in detail, HEF1 also localizes to focal adhesions in hTERT-RPE1 cells, as previously reported for many other cell lines. Shown, immunofluorescence of hTERT-RPE1 cells stained with antibody to HEF1 (green) and to acetylated α-tubulin (red) at 2 hours post-serum stimulation. Dotted line indicates HEF1 at focal adhesions; inset box shows basal body/centrosome. B. To confirm specificity of serum-indued phospho-AurA signal, antibody to T288-phospho-AurA (Cell Signaling) and total AurA was used to probe Western blots of untreated or phosphatase-treated hTERT-RPE1 cells at the times indicated after serum addition. CIP, calf intestinal phosphatase; PP1, protein phosphatase 1 (New England Biolabs) according to the manufacturer's recommendations.

FIG. 7. HDAC6 activity is necessary for resorption of cilia. A. Treatment of hTERT-RPE1 cells with histone deacetylase inhibitors prevents ciliary resorption. Cells were incubated with indicated compounds or vehicle (DMSO) at concentrations described in hereinbelow for 2 hours prior to induction of ciliary disassembly. The assay was performed 3 times, with an average of 100 cells counted/time point. B. TSA and tubacin increase intracellular levels of acetylated tubulin. Shown, Western blot with indicated antibodies showing levels of acetylated tubulin in cells treated with TSA, tubacin, niltubacin, or vehicle (DMSO) C. GSK3β inhibitor and farnesyltransferase inhibitor (FTI) do not inhibit ciliary disassembly. D. Depletion of HDAC6 restricts serum-induced disassembly of cilia in hTERT-RPE1 cells transfected for 48 hrs with siRNAs to HDAC6, HDAC2, or a scrambled control. E. Western analysis of hTERT-RPE1 cells treated with siRNA to HDAC6, HDAC2, or scrambled control. F. Active AurA or PBS were microinjected into hTERT-RPE1 cells pretreated for 2 hours with tubacin or DMSO.

FIG. 8. Depletion and FACS profiles of hTERT-RPE1 cells treated with siRNAs or small molecules. A. hTERT-RPE1 cells were treated with siRNA to AurA or HEF1, and protein depletion visualized by Western analysis. SiRNAs used were Ambion Cat.#44607 and 44668 for HEF1, and Dharmacon Cat.#D-003545-02 for AurA. B. Immunoprecipitates from experiments described in FIG. 3B were probed with antibody to AurA. C. FACS analysis of cells pre-depleted with siRNA to HEF1 or to AurA, or scrambled control (Scr) siRNA, at the times indicated after serum stimulation. D. Time course and degree of ciliary resorption assessed by scoring cells treated with siRNAs and inhibitors as indicated, based on visualization after staining with antibody to glutamylated tubulin. E. Cilia visualized with antibody to both acetylated α-tubulin (red) and glutamylated α-tubulin (blue), indicating equivalent staining. F. FACS analysis of cells pre-depleted with two siRNAs for HDAC6 (siHD6a, b), siRNA to HDAC2 (siHD2), or control scrambled (Scr) siRNA at the times indicated after serum stimulation. G. FACS analysis of cells pre-treated with DMSO vehicle, niltubacin, tubacin, or TSA, at the times indicated after serum stimulation.

FIG. 9. Direct phosphorylation by AurA activates HDAC6 tubulin deacetylase activity. A. hTERT-RPE1 whole cell lysate (WCL) of cells treated with AurA inhibitor PHA 680632 (+) or with vehicle (−) was analyzed by Western blot directly, or following immunoprecipitation (IP) with antibody to AurA, using antibodies as indicated. The immunoprecipitation of the slow-migrating form of HDAC6 is not impacted by treatment of cells with PHA-680632, indicating that it most likely represents HDAC6 modified by an additional (unknown) cellular kinase/s. B. AurA phosphorylates HDAC6. In vitro translated and immunoprecipitated HDAC2 or HDAC6 (HD2, HD6), or recombinant GST (−), were mixed with recombinant AurA and used in an in vitro kinase assay. Reaction was split and used for autoradiography (³²P) or Western Blot (WB) C. In vitro translated HDAC2 or HDAC6 (HD2, HD6) were immunoprecipitated (IPed). IPs were mixed with AurA (+) or buffer (−), then used for either an in vitro tubulin deacetylation assay, or in an in vitro kinase assay using γ-³²P-ATP. Reaction mix was visualized by Western blot and by autoradiography, as indicated. D. HDAC6 localizes to disassembling cilia 2 hours after serum treatment. Scale bar, 15 μM.

FIG. 10. Disassembly of cilia visualized with antibody to the ciliary marker glutamylated tubulin (1:250, Sigma). A. Time course and degree of ciliary resorption assessed by scoring cells treated with tubacin and niltubacin as indicated, based on visualization after staining with antibody to glutamylated tubulin. B. Experiment as in A., with cells microinjected with AurA, D274D, and GST. C. Staining with α-acetylated α-tubulin (red) and glutamylated tubulin (blue) indicates microinjection of AurA (marked by Dextran488, green) induces loss of signal at cilia during ciliary disassembly (boxes shown at high magnification to right; compare with uninjected cells, *). In contrast, AurA injection does not influence a-acetylation of cytoplasmic microtubule networks.

FIG. 11. A role for IFT proteins in AurA-induced ciliary resorption. A. Western blot demonstrating siRNA depletion of IFT88 (siIFT88) in ciliated hTERT-RPE1 cells at times following serum treatment, relative to scramble-depleted control. B. Immunofluorescence matching FIG. 11A at time 0, indicating relative degree of depletion of IFT88 at the basal body. C. Ciliary disassembly in IFT88-depleted (s88) versus Scr-depleted cells, at 0, 12, or 18 hours after serum treatment, based on the total cell population (gray bars). Black bars (right) indicate % ciliated cells at time 0 calculated specifically from cells confirmed by immunofluorescence to have significant IFT88 staining (88+), or to be well-depleted for IFT88 (88−). D. Cells treated with scrambled (Scr) or AurA-targeting (siAurA) siRNAs, or with PHA-680632 were fixed 2 hours after serum-initiated disassembly. Shown, immunofluorescence indicating cilia (anti-acetylated α-tubulin, red) and IFT88 (green). Insets are enlargements of boxed ciliary structures; arrows indicate direction of ciliary projection relative to basal body. Scale bars, 10 μm.

FIG. 12. Ciliary disassembly in IFT20- (s20) versus Scr-depleted cells.

FIG. 13. Working Model. A. Aurora A (AurA) and HEF1 are localized to the basal body of quiescent, ciliated cells. B. Our data are consistent with a model in which growth factors induce HEF1 expression, promoting HEF1-dependent activation of Aurora A. This results in phosphorylation of ciliary HDAC6 (H6) by Aurora A, thereby inducing ciliary resorption.

FIG. 14. Flow induced mechanosensation and diffusible factors promote signaling at cilia. Stimulation of poly-cystins 1 and 2 induces Ca²⁺ influx, activating Id2 and other effectors. Hedgehog (Hh) activation of its effector Gli requires Polaris (Pol) to anchor the Smoothened (Smo) receptor at the cilium. Similarly, PDGFaa interacts with its receptor at cilia to activate MEK and ERK, with contributions from Pak.

FIG. 15. Outline of crosses to generate mice for exploring efficacy of agents for the treatment of disorders related to aberrant cilia assembly and disassembly, particularly PKD. Progeny from F3 brother-sister matings will have the genotypes shown. *Only the genotypes of animals to be used in subsequent mating steps are shown.

FIG. 16. Calcium release rapidly induces AurA autophosphorylation. A. Immunofluorescence of HK-2 cells 30 seconds following stimulation with arginine vasopressin (+AVP), versus control untreated cells (−AVP). Cells were visualized with antibodies to AurA (red) and T288-phospho-AurA (phAurA, green), and DAPI (blue), as indicated; scale bar equals 20 μm. Insets show magnification of indicated centrosomes. The graph to right quantifies phAurA-positive cells at times after stimulation. An average of 150 cells were counted in each of three experiments, with error bars indicating the standard error (S.E.). *, p<0.05. B. Western analysis of AVP-treated HEK293 cells overexpressing AurA. β-actin is used in all Westerns as loading control. Note, endogenous levels of phAurA in HK2 cells are too low to visualize by Western blot. C. HEK293 cells overexpressing AurA were incubated with 5 μM thapsigargin for the indicated periods of time, and then cell lysates analyzed by Western blot. Graph below indicates ratio of phAurA to total AurA, quantified from analysis of blots; error bars show the S.E. *, p<0.05. D. AurA immunoprecipitation and in vitro kinase assay, following thapsigargin treatment. HEK293 cells overexpressing AurA were treated with 5 μM thapsigargin or control DMSO for 5 minutes, AurA immunoprecipitated, incubated with histone H3 substrate in an in vitro kinase assay. phHH3, antibody to phosphorylated histone H3. E. Experiment similar to that shown in D., except performed in the presence of 50 μM BAPTA/AM. F. AurA-overexpressing HEK293 cells transfected with siRNA depleting NEDD9 (siNEDD9) or scrambled control siRNA (scr), then were incubated with 5 μM thapsigargin and processed as in C. *, p<0.05.

FIG. 17. CaM binds and activates AurA. A. In vitro kinase using recombinant AurA, with the addition of CaM and Ca²⁺ as indicated, with MBP used as AurA substrate and histone H1 as negative control. ³²P-γ-ATP signal at ˜55 kDa indicates AurA autophosphorylation. B. In vitro kinase with purified AurA and CaM, CaM activates AurA autophosphorylation and phosphorylation of NEDD91-363 substrate. The AurA inhibitor PHA680632 (PHA) was used 1 μM. C. CaM Sepharose (CaM-S) was used for immunoprecipitation (IP) from whole cell lysates (WCL) transfected with plasmids expressing AurA-RFP and RFP, followed by Western blotting with antibodies indicated. D. AurA pulldown with CaM-Sepharose is Ca²⁺-dependent. Recombinant baculoviral AurA was supplemented with 1 mM CaCl₂ (Ca²⁺) or 1 mM EGTA and incubated with CaM-Sepharose (CaM-S) or control Sepharose (S). E. Lysates from HEK293 cells overexpressing AurA were preincubated for 1 hr with 10 μM of the CaM inhibitor Calmidazolium (CMZ) or control DMSO, incubated with 5 μM thapsigargin, and then analyzed by Western blot. Graph below indicates ratio of phAurA to total AurA, quantified from analysis of blots; error bars show the S.E. *, p<0.05.

FIG. 18. AurA and NEDD9 interact with PC2. A. Antibody to AurA or control IgM was used for IP from HK-2 cells, followed by Western blotting with antibodies to AurA or PC2 (G20), as indicated. Strong band migrating at ˜110 kDa represents native PC2, with faster migrating bands reflecting degradation products. B. Antibody to AurA was used for immunoprecipitation (IP) from whole cell lysates (WCL) transfected with plasmids expressing AurA, and either GFP-PC2₇₇₉₋₉₆₈ or GFP, followed by Western blotting with antibodies indicated. C. Antibody to AurA was used in IP from in vitro mixture containing GST-PC2₇₇₉₋₉₆₈ or GST only, and recombinant His-AurA. CB indicates Coomassie blue staining of starting material. D. RFP-fused catalytic (cat, aa 132-403) and non-catalytic (non-cat, aa 1-131) domains of AurA, or full length AurA-RFP, with co-expressed with FLAG-PC2₇₇₉₋₉₆₈ into HEK293 cells, and IPed with FLAG antibody, followed by Western blot with antibodies indicated. E. HEK293 cells were co-transfected with plasmids expressing combinations of GFP, GFP-PC2₇₇₉₋₉₆₈ or Flag-PC1-CT₄₁₉₁₋₄₃₀₂ and AurA, as indicated. Cell lysates were IPed using anti-GFP antibody, and visualized with antibodies indicated in Western blot analysis. F. Plasmids expressing HA-HEF1, HA-BioB, and GFP-PC2₇₇₉₋₉₆₈ were transfected into HEK293 cells, cell lysates IPed with antibody to GFP, and Western blotting performed with antibodies indicated.

FIG. 19. AurA phosphorylates PC2 at Ser-829. A. Structural motifs on PC2 C-terminal (CT domain) include PC1 binding motif (coiled-coil domain) (PC1-B a.a. 832-895), an EF-hand for Ca²⁺ binding (EF-hand, a.a. 754-782), and endoplasmic reticulum (ER) targeting sequences (ER-T, a.a. 787-820). Location of assessed sites for CK2 (S812) and AurA (S829, S944) phosphorylation are indicated. B. Bacterially expressed GST-PC2₇₇₉₋₉₆₈, or GST-NEDD₉₁₋₃₆₃ and MBP (positive controls), or histone H1 (H1) were incubated with recombinant active AurA kinase in an in vitro kinase assay with ³²P-γ-ATP. The reactions were resolved by SDS-PAGE, and analyzed by autoradiography, Coomassie blue (CB) staining, and Western blotting with antibody to AurA. C. Experiment as in B., except GST-NEDD₉₁₋₃₆₃ was included in reactions indicated. D. Bacterially expressed GST-PC2₇₇₉₋₉₆₈ was incubated with recombinant pre-activated AurA (rAurA) or AurA purified from baculovirus in an vitro kinase assay in the presence or absence of PC2, CaM, and Ca²⁺, as indicated. Top panel, autoradiography; lower panels, western blots with antibodies indicated. E., F. GST-fused derivatives of PC2₇₇₉₋₉₆₈, with mutations affecting a previously defined CK2 site (PC2S812A) and two candidate AurA sites (PC2S829A and PC2S944A) alone or in combination were used in in vitro kinase reactions with recombinant AurA. Phosphoimaging of autoradiographs from 3 independent experiments was quantified (F), and relative phosphorylation of PC2₇₇₉₋₉₆₈ mutants calculated (graph). Data are expressed as mean values±S.E. of three experiments. G. H. Experiments as in E and F., except recombinant CK2 was used rather than AurA. I. HEK293 cells were transfected with Myc-tagged PC2 or S829A mutant PC2 (Myc), in the presence or absence of AurA or T288D catalytically active AurA and then immunoprecipitated. Cells were treated with 10 μM H89 PKA inhibitor (Calbiochem). or 500 nM PHA680632 (PHA) AurA inhibitor for 3 hr as indicated. RRxS-directed antibody was used to visualize phosphorylation of the S829A residue. Western blot of coimmunoprecipitation from 3 independent experiments was quantified (J), and relative phosphorylation of PC2 calculated (graph). *, significant difference versus PC2 (p<0.05); ** difference from PC2 is not significant.

FIG. 20. AurA binds PC2 and negatively regulates PC2-dependent Ca²⁺ release. A. HEK293 cells transiently cotransfected 48 hours previously with plasmids expressing PC2, and AurA-RFP or control RFP, were loaded with 5 uM Fluo-4 AM, which auto-fluoresces upon binding cytoplasmic Ca²⁺. Immunofluorescence was measured before and after addition of AVP (indicated by arrow). Data is presented as F/F0 ratio, indicating increase in signal over unstimulated cells. B. For this and subsequent experiments, the mean increase in amplitude (±S.E.) of AVP-induced cytoplasmic calcium transients, was calculated from n>50 RFP-positive cells in each of 3 experiments. *, AurA-RFP amplitude was decreased versus RFP controls, p=0.008. C. HEK293 cells transiently transfected 48 hours previously with a plasmid expressing PC2 or vector control, and pretreated 2 hours with 500 nM PHA 680632 or DMSO vehicle, were analyzed as in A. D. Quantification of data in C. *, PHA 680632 increased amplitude of signal versus DMSO, p=0.0007. E. HK2 cells stably expressing PC2 or vector control, pretreated for 2 hours with 500 nM PHA 680632 or DMSO vehicle, were processed as in A. F. Quantification of data in E. PHA 680632 increased amplitude of signal versus DMSO in PC2 transfected cells (*,p=0.00065) and in untransfected HK-2 cells (**, p=0.009). G., H. siRNA depletion of AurA (siAurA) in HK-2 cells increases amplitude of calcium release versus scrambled control (Scr). H. Quantification shows significant difference in amplitude (*, p=0.0055).

FIG. 21. A cytoplasmic pool of AurA regulates ER-localized PC2. A. Immunofluorescence of HK2 cells treated with siRNA to deplete AurA (siAurA), or with control scrambled siRNA (Scr). Scale bar equals 10 μm. B. AVP-induced increase in cytoplasmic Ca²⁺ in ciliated versus non-ciliated HK2 cells stably expressing PC2 that were pretreated for 2 hours with PHA 680632 or DMSO vehicle. Graph to right compares maximum amplitude of signal. Differences in calcium release between ciliated and unciliated cells were not significant. *, p<0.05. C. AurA-dependent phosphorylation does not play a role in subcellular localization of PC-2. Exogenous PC-2 stably expressed in ciliated and non-ciliated HK2 cells (left) and endogenous PC2 in HK2 cells (right) after PHA-680632 or DMSO treatment are comparably sensitive to Endo H treatment, consistent with location in ER and pre-middle Golgi membrane compartments in HK2 cells. D. Immunofluorescence localization of wild type PC2, or PC2S829E and PC2S829A, in relation to markers for the endoplasmic reticulum (PDI, red) or nucleus (DAPI, blue), scale bar equals 10 μm. E. endo H mapping of mutants. Assay as in D., except comparing PC2±AurA, or PC2S829E and PC2S829A. In right two lanes, cells have been transfected with plasmid overexpressing AurA.

FIG. 22. Abundant AurA in human kidneys and cysts provides a potential therapeutic target for PKD. A. Immunohistochemical (IHC) visualization of endogenous AurA and phAurA in normal human kidney tissue. Typical fields from cortical (left) and medullary region (center) are shown. Right, sections stained with blocking peptide preincubated with antibody to total AurA and negative control IgG. PCT, proximal convoluted tubules; DCT, distal convoluted tubules; CD, collecting ducts; LH, thin segments of Loops of Henle; GM, glomerulus. B. IHC detection of AurA and phAurA in renal cysts from PKD patients. Epithelial cells lining cysts, but not cyst-associated fibrotic tissue, consistently demonstrate positive staining. C. IC₅₀ curve for PHA-680632 (PHA) treatment of HK2 cells stably expressing PC2. Cell viability was detected by Alamar blue assay 72 hours post-treatment. D. HK2 cells stably expressing PC2 were treated with DMSO or with PHA-680632 (PHA) at concentrations of 0.5 or 3.25 μM for 2 hours (2 h) or 24 hours (24 h), prior to AVP stimulation and determination of F/F0 ratio. E. Quantification of relative F/F0 amplitude of data in D. Difference between PC2/DMSO and * is significant (P<0.01); difference between ** and * is significant, P=0.0022; difference between * and * is not significant.

FIG. 23. Schematic diagram of the role played by Aurora A in the regulation of cellular homeostasis for calcium.

DETAILED DESCRIPTION OF THE INVENTION

The mammalian cilium protrudes from the apical/lumenal surface of polarized cells, and acts as a sensor of environmental cues. Numerous developmental disorders and pathological conditions have been shown to arise from defects in cilia-associated signaling proteins. Despite mounting evidence that cilia are essential sites for coordination of cell signaling, little is known about the cellular mechanisms controlling their formation and disassembly. Here we show that defined interactions between the pro-metastatic scaffolding protein HEF1/Cas-L/NEDD9 and the oncogenic Aurora A (AurA) kinase at the basal body of cilia causes phosphorylation and activation of HDAC6, a tubulin deacetylase, promoting ciliary disassembly. We show that this pathway is both necessary and sufficient for ciliary resorption, and constitutes an unexpected non-mitotic activity of AurA in vertebrates. Moreover, we demonstrate that small molecule inhibitors of AurA and HDAC6 selectively stabilize cilia from regulated resorption cues, suggesting a novel mode of action for these clinical agents.

Oncogenic hyperactivation of the mitotic kinase Aurora-A (AurA) in cancer is associated with genomic instability. Increasing evidence indicates AurA also regulates critical processes in normal interphase cells, but the source of such activity has been obscure. We report that multiple stimuli causing release of Ca2+ from intracellular endoplasmic reticulum (ER) stores activate AurA by inducing direct Ca2+-dependent calmodulin binding to AurA. Subsequently, activated AurA binds, phosphorylates, and limits the activity of the Ca2+-permeable nonselective cation channel polycystin 2 (PC2/TRPP2, encoded by the polycystic kidney disease-associated gene PKD2), limiting the amplitude of Ca2+ release from the ER. Active AurA is abundant in non-mitotic cells in normal kidneys and elevated in cells lining PKD-associated renal cysts, and inhibitors of AurA significantly enhance PC2-dependent Ca2+-release. These and other findings provide a new context for evaluating AurA function in normal cells and cancer, and suggest AurA may be a relevant new modifier gene for PKD.

The following definitions are provided to facilitate an understanding of the present invention:

Disorders associated with aberrant cilia function and regulation include, without limitation, polycystic kidney disease (PKD), Bardet-Biedl Syndrome (BBS), renal cysts, infertility, respiratory disorders, situs inversus, and predisposition to obesity, diabetes, and hypertension.

The phrase “aurora kinase inhibitor” refers to any agent which functions to inhibit or down regulate aurora kinase A and/or aurora kinase B. Such agents include, without limitation, small molecules, chemical compounds and nucleic acid molecules which function to down regulate expression of target genes. Exemplary agents include C1368, PHA-680632, hesparadin, MLN 8327, VX-680, MLN8054, and siRNA which hybridize selectively to aurora kinase encoding mRNA and down regulate expression of the aurora kinase protein product. Exemplary siRNAs that target aurora kinase have the following sequence: Hs_AURKA_(—)1 TCCCAGCGCATTCCTTTGCAA and Hs_STK6_(—)5 CACCTTCGGCATCCTAATATT.

The terms “transform”, “transfect”, “transduce”, shall refer to any method or means by which a nucleic acid is introduced into a cell or host organism and may be used interchangeably to convey the same meaning. Such methods include, but are not limited to, transfection, electroporation, microinjection, PEG-fusion and the like. The introduced nucleic acid may or may not be integrated (covalently linked) into nucleic acid of the recipient cell or organism. In bacterial, yeast, plant and mammalian cells, for example, the introduced nucleic acid may be maintained as an episomal element or independent replicon such as a plasmid. Alternatively, the introduced nucleic acid may become integrated into the nucleic acid of the recipient cell or organism and be stably maintained in that cell or organism and further passed on or inherited to progeny cells or organisms of the recipient cell or organism. Finally, the introduced nucleic acid may exist in the recipient cell or host organism only transiently.

The term “selectable marker gene” refers to a gene that when expressed confers a selectable phenotype, such as antibiotic resistance, on a transformed cell.

The term “operably linked” means that the regulatory sequences necessary for expression of the coding sequence are placed in the DNA molecule in the appropriate positions relative to the coding sequence so as to effect expression of the coding sequence. This same definition is sometimes applied to the arrangement of transcription units and other transcription control elements (e.g. enhancers) in an expression vector.

“Native” refers to a naturally occurring (“wild-type”) nucleic acid sequence.

“Heterologous” sequence refers to a sequence which originates from a foreign source or species or, if from the same source, is modified from its original form.

A “coding sequence” or “coding region” refers to a nucleic acid molecule having sequence information necessary to produce a gene product, when the sequence is expressed.

The term “animal” is used herein to include all vertebrate animals, except humans. It also includes an individual animal in all stages of development, including embryonic and fetal stages. A “transgenic animal” is any animal containing one or more cells bearing genetic information altered or received, directly or indirectly, by deliberate genetic manipulation at the subcellular level, such as by targeted recombination or microinjection or infection with recombinant virus. The term “transgenic animal” is not meant to encompass classical cross-breeding or in vitro fertilization, but rather is meant to encompass animals in which one or more cells are altered by or receive a recombinant DNA molecule. This molecule may be specifically targeted to defined genetic locus, be randomly integrated within a chromosome, or it may be extrachromosomally replicating DNA. The term “germ cell line transgenic animal” refers to a transgenic animal in which the genetic alteration or genetic information was introduced into a germ line cell, thereby conferring the ability to transfer the genetic information to offspring. If such offspring in fact, possess some or all of that alteration or genetic information, then they, too, are transgenic animals.

The alteration or genetic information may be foreign to the species of animal to which the recipient belongs, or foreign only to the particular individual recipient, or may be genetic information already possessed by the recipient. In the last case, the altered or introduced gene may be expressed differently than the native gene.

The DNA used for altering a target gene may be obtained by a wide variety of techniques that include, but are not limited to, isolation from genomic sources, preparation of cDNAs from isolated mRNA templates, direct synthesis, or a combination thereof.

Methods for Treating PKD

As explained above, the present invention is directed towards methods for modulating or alleviating the symptoms of polycystic kidney disease comprising, the administration of an Aur-A inhibitor such as those listed above. Administration can be achieved by any suitable route, such as parenterally, transmucosally, e.g., orally, nasally, or rectally, or transdermally. Preferably, administration is parenteral, e.g., via intravenous injection. Alternative means of administration also include, but are not limited to, intra-arteriole, intramuscular, intradermal, subcutaneous, intraperitoneal, intraventricular, and intracranial administration, or by injection into the renal cyst being treated.

The Aur-A inhibitor may be employed in any suitable pharmaceutical formulation, as described above, including in a vesicle, such as a liposome [see Langer, Science 249:1527-1533 (1990); Treat et al., in Liposomes in the Therapy of Infectious Disease and Cancer, Lopez-Berestein and Fidler (eds.), Liss: New York, pp. 317-327, see generally, ibid] Preferably, administration of liposomes containing the agents of the invention is parenteral, e.g., via intravenous injection, but also may include, without limitation, intra-arteriole, intramuscular, intradermal, subcutaneous, intraperitoneal, intraventricular, and intracranial administration, or by injection into the renal cyst being treated.

In yet another embodiment, a pharmaceutical composition of the present invention can be delivered in a controlled release system, such as using an intravenous infusion, an implantable osmotic pump, a transdermal patch, liposomes, or other modes of administration. In a particular embodiment, a pump may be used [see Langer, supra; Sefton, CRC Crit. Ref. Biomed. Eng. 14:201 (1987); Buchwald et al., Surgery 88:507 (1980); Saudek et al., N. Engl. J. Med. 321:574 (1989)]. In another embodiment, polymeric materials can be used [see Medical Applications of Controlled Release, Langer and Wise (eds.), CRC Press: Boca Raton, Fla. (1974); Controlled Drug Bioavailability, Drug Product Design and Performance, Smolen and Ball (eds.), Wiley: New York (1984); Ranger and Peppas, J. Macromol. Sci. Rev. Macromol. Chem. 23:61 (1983); see also Levy et al., Science 228:190 (1985); During et al., Ann. Neurol. 25:351 (1989); Howard et al., J. Neurosurg. 71:105 (1989)]. In yet another embodiment, a controlled release system can be placed in proximity of the target tissues of the animal, thus requiring only a fraction of the systemic dose [see, e.g., Goodson, in Medical Applications of Controlled Release, supra, vol. 2, pp. 115-138 (1984)]. In particular, a controlled release device can be introduced into an animal in proximity of the site of inappropriate immune activation or a tumor. Other controlled release systems are discussed in the review by Langer [Science 249:1527-1533 (1990)].

The examples set forth below are provided to exemplify certain embodiments of the invention. They are not intended to limit the invention in any way.

Example I

We demonstrate that an association between AurA and HEF1 at cilia in response to extracellular cues is required for ciliary disassembly. We also show that AurA activation is independently sufficient to induce rapid ciliary resorption, and that AurA acts in this process through phosphorylating HDAC6, thus stimulating HDAC6-dependent tubulin deacetylation (Hubbert et al., 2002) and destabilizing the ciliary axoneme. Importantly, our identification of a spatiotemporally restricted action of AurA at the ciliary basal body in cells emerging from G0 demonstrates an unexpected non-mitotic activity for AurA in vertebrate cells. We also determine that small molecule inhibitors of AurA and HDAC6 reduce regulated disassembly of cilia, which may have important implications for the action of these drugs in the clinic. Together, these data reveal important activities for HEF1, AurA, and HDAC6 in regulation of ciliary resorption, which should also inform the actions of these proteins in cell cycle and cancer (Hideshima et al., 2005; Kim et al., 2006; Marumoto et al., 2005; Pugacheva and Golemis, 2005).

The following materials and methods are provided to facilitate the practice of the present example.

Cell culture and siRNA. hTERT-RPE1 cells were grown in DMEM with 10% fetal bovine serum (FBS). For analysis of ciliary disassembly, cells were plated at 30% confluence in plates containing glass cover slips, and starved for 48 hours (in Opti-MEM or regular DMEM, without added serum) to induce cilia formation, followed by treatments described hereinbelow. For siRNA treatment, cells were initially plated in DMEM/10% FBS in plates containing cover slips, and 12 hours later siRNA transfection was performed in Opti-MEM with Oligofectamine (Invitrogen) according to manufacturer recommendations, and fixed 48 hours after transfection, following treatments indicated in Results. The remaining cells on plate were lysed, then either directly analyzed by Western blot analysis, or used for immunoprecipitation (IP)-kinase reaction to measure AurA activity.

For RNA interference (RNAi)-induced depletion of HEF1 and AurA, 2 independent, synthetic duplex siRNAs were used for each gene: 1) Ambion, cat#16704, NEDD9, ID:17729. sense 5′GGUAUAUCAGGUGCCACCAtt3′; 2) Dharmacon, custom, sense: 5′ AAGGGGUAUAUGCCAUUCCGCdTd T 3′⁵⁷. Non-specific control siRNAs including scrambled (Dharmacon, cat#D-001206-13-05) and GFP-directed sequences (Dharmacon, cat#D-001300-01-20) were used for reference.

Drug inhibition experiments. The Aurora kinase inhibitor PHA-680632, GSK3β-inhibitor 1 (Calbiochem), FTI-277 (Calbiochem), Tubacin, Niltubacin or DMSO vehicle were added to hTERT-RPE1 cells 2 hours prior to the initiation of ciliary disassembly. After initial titration experiments to establish effective range, PHA-680632 was used at 0.5 μM, Tubacin and Niltubacin at 2 μM, GSK3β-inhibitor 1 at 2 μM, FTI-277 at 50 nM concentration for the experiments described. Protein expression, Western blotting, and immunoprecipitation. For microinjection, recombinant glutathione-S-transferase (GST), GST fused AurA mutants T288A and D274N) produced from BL21 (DE3) bacteria were purified using the MicroSpin GST Purification Module (Amersham Biotech.). Purified recombinant AurA was purchased from Upstate; this AurA was pre-activated based on incubation with ATP. Mutationally inactive AurA (T288A,) was also made using a baculoviral expression system (Invitrogen), and was purified by Ni-Sepharose 6FF (Amersham). To prepare lysates for Western blotting and IP, mammalian cells were disrupted by M-PER lysis buffer (Pierce) supplemented with EDTA-free protease inhibitor cocktail (Roche). Lysates used for IP were incubated overnight with antibody at 4° C., subsequently incubated for 2 hours with protein A/G-sepharose (Pierce), washed, and resolved by SDS-PAGE. Western blotting was performed using standard procedures and proteins visualized using the West-Pico system (Pierce). Antibodies used included mouse monoclonal antibody (mAb) anti-HEF1 2G9 (Pugacheva and Golemis, 2005), anti-α-tubulin mAb (Sigma), anti-AurA (BD Bioscience) for Western blotting, anti-AurA rabbit polyclonal (Cell Signaling) for IP, anti-Phospho-AurA/T288 (BioLegend), anti-Phospho-AurA/T288 (Cell Signaling), anti-HDAC6 rabbit polyclonal (Upstate; 1:5000), anti-HDAC2 rabbit polyclonal (Invitrogen) and mAb anti-β-actin (AC15, Sigma), anti-IFT88 and anti-IFT20. Secondary horseradish peroxidase (HRP)-conjugated antibodies were from Amersham Biotech. Immunofluorescence. Cells were fixed with 4% paraformaldehyde (10 min) then methanol (5 min), permeabilized with 1% Triton-X100 in PBS, blocked in 1×PBS, 3% BSA, and incubated with antibodies using standard protocols. Primary antibodies included rabbit polyclonal anti-Aurora A and anti-phospho-AuroraA/T288, (Cell Signaling), mouse mAb anti-HEF1 (14A11), polyclonal anti-γ-tubulin (Sigma), anti-α-tubulin mAb (Sigma), anti-acetylated α-tubulin mAb (clone 6-11B-1, Sigma, and clone K(Ac)40 Biomol), anti-IFT88 and anti-IFT20 (gifts of G. Pazour), mouse anti-glutamylated tubulin (Sigma), and anti-HDAC6 (Upstate). Secondary antibodies labeled with Alexa-488, Alexa-568, and Alexa-633, and TOTO-3 dye to stain DNA, were from Molecular Probes/Invitrogen. DNA was co-stained in some experiments by propidium iodine (Sigma) or Draq5 (Alexis). Confocal microscopy was performed using a Radiance 2000 laser scanning confocal microscope ((Carl Zeiss, Thornwood, N.Y.) coupled to a Nikon Eclipse E800 upright microscope (Nikon). Statistical analysis of data by one-way ANOVA was performed using GraphPad Instat 3.0 (San Diego, Calif.). Microinjection. Microinjections were performed on a Nikon TE300 Microscope (Nikon, Melville, N.Y.) that was equipped with an Eppendorf Transjector 5246 semi-automatic microinjector and micromanipulator (Eppendorf, Westbury, N.Y.). Cells were plated on gridded coverslips (Belco) and starved for 48 hours before cytoplasmic microinjection of 0.05 μM pre-activated AurA (Upstate), inactive AurA (T288A) and (D274N), GST protein, or buffer. Proteins were pre-filtered through a 0.2-μm Milliopore membrane and mixed with Dextran Green488 (Molecular Probes) to mark injected cells. Injected cells were incubated at 37° C. before fixation. Typically, 150 cells were microinjected in each of 3 experiments. Kinase and tubulin deacetylation assays. In vitro kinase assays were performed using recombinant active AurA (Upstate), mutationally inactive AurA purified from baculovirus and BL21 (DE3) bacteria, or endogenous AurA immunoprecipitated from mammalian cells. A standard kinase reaction with γ-³²P(ATP) and histone H3 and MBP (Upstate) substrates was done as in (Pugacheva and Golemis, 2005). For deacetylase assays, HDAC6 and HDAC2 were in vitro translated using a TnT-Coupled Reticulocyte Lysate System (Promega), immunoprecipitated, and incubated with/without active AurA(Upstate) in the presence of (25 μg) stabilized microtubules prepared from purified bovine brain tubulin (Cytoskeleton) to measure deacetylase activity (as in (Hubbert et al., 2002)) and with γ-³²P-ATP (Perkin-Elmer) in AurA reaction buffer. 1/10 volume of samples were reserved for Western blotting.

Results A System for Regulated Ciliary Assembly and Disassembly

We established a system to study ciliary dynamics in the hTERT-RPE1 cell line. 48 hours after plating cells at 50-70% confluence in Opti-MEM medium without serum, >80% of hTERT-RPE1 cells had clearly visible cilia (FIG. 1A). Cilia were typically of 3-4 μm length, with an acetylated α-tubulin-marked axoneme adjacent to two γ-tubulin-positive structures reflecting the basal body and the second cellular centriole. See FIG. 2A. Treatment of these ciliated cells with medium containing 10% fetal bovine serum (FBS) caused ciliary disassembly over the following 24 hours (FIG. 1B). This disassembly occurred in two waves, with the first occurring 1-2 hours after serum stimulation, and the second after 18-24 hours. FACS analysis, BrDU staining, and observation of condensed DNA and mitotic figures indicated that cells remained predominantly in G1 phase at 2 hours post-serum addition, while during the 18-24 hour disassembly wave, most cells were entering mitosis (FIGS. 2B, 2C). This disassembly behavior was not unique to hTERT-RPE1 cells, as we observed a comparable biphasic resorption profile in the IMCD-3 murine and Caki-1 human renal cell lines (FIGS. 2D, 2E). To begin to assess serum components that might regulate ciliary disassembly, we have assessed PDGF, TGF-β, and EGF (FIG. 4). Of these, only PDGF elicited a partial response. Full disassembly likely requires the combined input of several distinct serum components.

Dynamic regulation of HEF1 and AurA at the basal body during ciliary disassembly. AurA (FIG. 1C) and HEF1 (FIGS. 1D,E) localized to the basal body and the second centriole in quiescent, ciliated hTERT-RPE1 cells. In contrast, activated (T²⁸⁸-phosphorylated) AurA was not detected at basal bodies of cilia in quiescent cells (FIG. 1F, 1I (0 h)) under fixation conditions at which it was clearly evident in mitotic cells (FIG. 1G).

If AurA were functionally important for ciliary disassembly, we would expect changes in the activity of AurA 1-2 hours after serum treatment, potentially accompanied by changes in the AurA activator HEF1. Indeed, HEF1 expression increased at 1-2 hours after serum stimulation, dropped, and peaked again at ˜18-24 hours after serum stimulation (FIG. 1H). HEF1 initially appeared as a faster migrating 105 kD species, with a slower migrating 115 kD species appearing later. This 115 kD species represents S/T-phosphorylated HEF1, is most abundant during the G2/M compartment in actively cycling cells, and is associated with AurA activation (Law et al., 1998; Pugacheva and Golemis, 2005). Total AurA levels sometimes increased slightly at 2 hours post-serum stimulation, but were largely unaffected (FIG. 1H). In contrast, peaks of phospho-T²⁸⁸-AurA appeared precisely at each of the two waves of ciliary disassembly (FIGS. 1H, I). Strikingly, phospho-T²⁸⁸-AurA was almost never detected at a basal body near a well-formed cilium. Although phospho-T²⁸⁸-AurA invariably co-localized with both γ-tubulin-marked basal bodies/centrioles and with total AurA, in 85-90% of cells with phospho-T²⁸⁸-AurA, centrioles had no accompanying cilium. In 10-15% of cells with phospho-T²⁸⁸-AurA, centrioles with adjacent acetylated α-tubulin-marked cilia were observed, but these cilia were significantly shortened (˜1-2 versus 3.5 μm) (FIG. 1I). Similar profiles of HEF1 and AurA expression and activation were observed in serum-treated IMCD3 and Caki-1 cells, and PDGF-treated hTERT-RPE1 cells (FIGS. 3B, 3C, 3F, 3G). The simplest interpretation of these results is that activation of AurA at the basal body immediately precedes the rapid disassembly of cilia.

HEF1-dependent activation of AurA induces ciliary disassembly. We used two complementary approaches to establish that AurA activation is necessary and sufficient for induction of ciliary disassembly, and that HEF1 is likely to contribute to this process.

First, exponentially growing hTERT-RPE1 cells were treated with siRNA targeting AurA or HEF1, or with control siRNA, plated for 2 days in OptiMEM to allow cilia formation, then treated with serum to induce ciliary disassembly. Immunoblotting confirmed siRNA treatment efficiently depleted AurA and HEF1 (FIG. 8A). AurA depletion blocked and HEF1 depletion greatly limited serum-induced disassembly (FIG. 3A). AurA activation was substantially reduced in cells treated with siRNA to HEF1 (FIG. 3B); this correlated with reduced levels of AurA in HEF1-depleted cells (FIG. 8B), implying HEF1 contributes to AurA stabilization as well as activation. Particularly at the second wave of ciliary disassembly, the residual cilia in HEF1-depleted cells were significantly longer than those in control cells (FIG. 3C), implying that HEF1 modulates the disassembly process. Importantly, cells treated with siRNA to AurA or HEF1, or with control siRNA, were all ˜80% ciliated before addition of serum, leading us to conclude that the predominant role for HEF1 and AurA is at the time of disassembly, i.e., these proteins are not required to form cilia.

Second, we used the small molecule AurA kinase inhibitor PHA-680632 (Nerviano Medical Sciences (Soncini et al., 2006)) to inactivate AurA kinase (FIGS. 3D,E). Disassembly of cilia was strongly reduced in cells pre-treated for 3 hours with 500 nM PHA-680632 (FIG. 3D). Although some ciliary disassembly was observed at 1 and 2 hours after serum stimulation, the percentage was lower than in DMSO-treated cells, and disassembly was not maintained, with cilia consistently re-established at the 8- and 12-hour time points. The second wave of ciliary disassembly, at the time of mitosis, was completely eliminated in PHA-680632-treated cells (FIG. 3D). In cells with inhibited AurA, hyper-phosphorylated HEF1 did not accumulate significantly at either wave of ciliary disassembly, indicating AurA dependence of this phosphorylation (FIG. 3E). Western blot (FIGS. 3E, 3F (right panels)), in vitro kinase assays (FIG. 3F, left panels) and immunofluorescence (FIG. 3G) confirmed the effectiveness of the compound in blocking AurA activation.

Together, these data imply that activation of AurA by HEF1 contributes to resorption of cilia at 2 and 18 hours following serum stimulation (FIG. 3A-E) and that active AurA is necessary to stably complete the disassembly process, but that HEF1 may not be the sole factor driving AurA activation and ciliary resorption (FIG. 3A). Further, FACS analysis of cells with siRNA-depleted HEF1 or AurA (FIG. 8C), or drug-inhibited AurA (FIG. 3H) AurA indicated that the blocked resorption of cilia at the 2 h time point does not reflect an indirect consequence of altered cell cycle compartmentalization due to AurA inhibition. Cells indeed show predictable siRNA- and drug-induced accumulation in G2 at 18-24 h after serum stimulation, which may account for the reduced resorption at these time points. However all cells at 2 h post-serum treatment have similar cell cycle profiles, remaining predominantly in G0/G1. Hence, the role of HEF1 and AurA at this early non-mitotic time point represents an unexpected direct action of these proteins.

AurA activation is sufficient to induce rapid disassembly of cilia. Next, as a direct approach to establish sufficiency of active AurA to induce disassembly, we microinjected pre-activated wild type AurA (aAurA), T288A AurA (a hypomorphic mutant, (Satinover et al., 2004)), D274N AurA (an inactive mutant), GST, or buffer alone, together with fluorescent marker dye, into hTERT-RPE1 cells with pre-formed cilia. Microinjection of aAurA rapidly induced the disappearance of cilia from cells maintained in low serum medium: Essentially as soon as cells could be fixed after microinjection, more than 80% of injected cells lacked cilia (FIGS. 5A, 5B). In contrast, injection of GST or buffer did not induce loss of cilia. Of the two mutants, D274N did not induce loss of cilia, while T288A caused eventual partial loss of cilia (FIG. 5A) and ciliary shortening (results not shown). The ability of aAurA, T288A, and D274N paralleled the behavior of these proteins in in vitro kinase assays performed in parallel to microinjections (FIG. 5C). Whereas aAurA was highly active and D274N was completely inactive, T288A became weakly active following brief incubation with cell lysates. Hence, the delayed resorption of cilia and ciliary shortening induced by T288A likely reflects the gradual emergence of an active pool of AurA following microinjection. HDAC6 is required for ciliary disassembly. Little is known about the cellular machinery necessary for disassembling cilia. In seeking targets of AurA phosphorylation that might be relevant to this process, we considered the possibility that the acetylated α-tubulin commonly used to visualize cilia might play an active role in stabilizing the ciliary axoneme, based on reports that α-tubulin deacetylation promoted the in vivo destabilization of microtubules (Matsuyama et al., 2002). In particular, histone deacetylase 6 (HDAC6) has been identified as an important cytoplasmic tubulin deacetylase that influences mitosis and chemotaxis through regulating tubulin stability (Hubbert et al., 2002).

To assess whether altered regulation of tubulin acetylation might mediate HEF1/AurA signaling, we treated ciliated hTERT-RPE1 cells with small molecule deacetylase inhibitors, and established the ciliary disassembly profile (FIG. 7A). Both the broad-spectrum HDAC inhibitor trichostatin A (TSA), and tubacin, an inhibitor specifically targeting HDAC6 (Hideshima et al., 2005), completely blocked serum-induced ciliary disassembly, whereas niltubacin, an inactive analog of tubacin, and vehicle alone had no effect. Levels of acetylated tubulin were measured in treated cells, confirming that these were increased in cells treated with TSA and tubacin, but not in cells treated with niltubacin or control vehicle (FIG. 7B). As a control, because both AurA and HDAC inhibitors blocked ciliary disassembly, we considered the possibility that regulated ciliary disassembly might be generally sensitive to signaling inhibitors because of non-specific toxicities. However, serum induced disassembly with a normal profile in cells treated with inhibitors of GSK-3β and farnesyltransferase (FTI), indicating that blocked ciliary disassembly was specific response to impaired AurA and HDAC6 signaling (FIG. 7C).

To further confirm a specific requirement for HDAC6, we next established that cilia do not disassemble in serum-treated cells with siRNA-depleted HDAC6 (FIG. 7D, 7E). Finally, we have microinjected aAurA into ciliated cells pre-treated for 3 hours with tubacin (FIG. 7F). Tubacin pre-treatment substantially limited the ability of microinjected AurA to disassemble cilia. Initial disassembly was slower, and in some cases transient, with a significant percentage of injected cells re-forming cilia by 1 hour after injection. As for AurA, neither tubacin treatment nor siRNA to HDAC6 influenced cell cycle profile at 2 h after serum stimulation, although both treatments led to accumulation in G2 at the later time point (FIGS. 8F, 8G). As a final control, we again used antibody to glutamylated tubulin as an independent means of scoring ciliary disassembly (FIG. 8E). The results of these experiments are equivalent to those obtained using antibody to acetylated α-tubulin (FIGS. 10A-C). Based on these data, we concluded that HDAC6 is an important downstream AurA effector for ciliary disassembly.

AurA phosphorylates HDAC6 to activate tubulin deacetylase activity. Taken together, our data suggested that the mechanism of ciliary disassembly by AurA requires intact HDAC6 deacetylation activity, to destabilize microtubules. AurA-dependent regulation of tubulin deacetylation may be direct or indirect. Importantly, although microinjection of AurA induced loss of ciliary α-acetylated tubulin as cilia disassemble, the non-ciliary α-acetylation of cytoplasmic microtubule networks were unaffected, suggesting a specific action of AurA and HDAC6 at the cilia (FIG. 10C). Further supporting this idea, HDAC6 localized to cilia in serum-starved cells and during the ciliary disassembly process (FIG. 9D and unpublished results), providing a ready target for AurA phosphorylation. Demonstrating a direct AurA-HDAC6 connection, antibody to AurA coimmunoprecipitated HDAC6 from hTERT-RPE1 cells (FIG. 9A). AurA-HDAC6 coimmunoprecipitation was not eliminated by pre-treatment of cells with PHA-680632, indicating that the association was not regulated by AurA activation status (FIG. 9A).

To directly determine whether HDAC6 might be an AurA substrate, recombinant activated AurA was used in an in vitro kinase assay with purified HDAC6, HDAC2, or GST, as in (Pugacheva and Golemis, 2005). AurA phosphorylated HDAC6, but not HDAC2 or the GST negative control (FIG. 9B). We next immunoprecipitated in vitro translated HDAC6 and a negative control, HDAC2, and gauged the relative ability of AurA to phosphorylate these proteins, and stimulate a tubulin deacetylase activity, in a defined in vitro assay. In reactions containing comparable levels of HDAC2 and HDAC6, only HDAC6 was phosphorylated by AurA (FIG. 9C). Moreover, AurA-phosphorylated HDAC6 was much more potent than unphosphorylated HDAC6 in deacetylating α-tubulin (FIG. 9C). These results lead us to conclude that AurA phosphorylation of HDAC6 stimulates HDAC6 deacetylase activity.

Ciliary disassembly and intraflagellar transport (IFT). Intraflagellar transport proteins perform important roles in mediating transport of proteins to and from the apical tip of cilia, and in many cases mutations in IFT proteins have been linked to ciliary dysfunction, loss of cilia, and pathological conditions (Sloboda, 2005). In contrast to depletion of HEF1 or AurA, depletion of representative IFT proteins IFT88 (FIGS. 6A-C) and IFT20 (FIG. 12) limits the initial formation of cilia in hTERT-RPE1 cells similar to reports in other cell types (Follit et al., 2006; Pazour et al., 2000). Based on immunofluorescence, cilia were only observed in IFT-depleted cells that retain at least some detectable IFT protein (FIG. 11C). This clear requirement of IFT proteins for ciliary assembly hinders the dissection of the contribution of these proteins in disassembly. However, intriguingly, the existing cilia in IFT88- or IFT20-depleted cells undergo minimal disassembly following serum stimulation, with the difference particularly noticeable at the early (2 h) timepoint (FIGS. 11C, 12). Further, depletion or inhibition of AurA alters the localization of IFT88 during the ciliary disassembly process. In untreated cells, IFT88 is seen intensely at the basal body and more diffusely along the axoneme of residual cilia two hours after serum stimulation, whereas in cells lacking active AurA, IFT88 accumulates at both the basal body and apical tip at this time point (FIG. 11D). It is likely that as in Chlamydomonas (Pan and Snell, 2005), IFT signaling mediates some aspects of ciliary disassembly.

Discussion

Cilia and flagella have been described as cellular “antennas”, sensing a multiplicity of extracellular stimuli to induce an intracellular response (Singla and Reiter, 2006). In addition to undergoing regulated resorption induced by extracellular cues, for over four decades cilia have been known to be dynamically resorbed and resynthesized throughout the cell cycle. Taken in sum, our data suggest a model (FIG. 13) in which the serum growth factor-induced activation of a HEF1-AurA complex allows AurA to phosphorylate and activate HDAC6, which destabilizes the ciliary axoneme by deacetylating tubulin. Unexpectedly, activation of AurA is a central component of this cascade even during the G1 resorption wave, indicating a non-mitotic activity for AurA in animals.

An important finding of this work is the novel connection between AurA and HDAC6. HDAC6 tightly interacts with α and β tubulins through its HDAC domain, which may restrict its enzymatic activity, based on reports that taxol treatment causes HDAC6 to accumulate on microtubules, and is accompanied by increased tubulin acetylation (Zhang et al., 2003). Localized phosphorylation by AurA may increase the turnover of HDAC6 at microtubules, thus increasing the active pool of HDAC6 at cilia. Interestingly, studies in Chlamydomonas indicate that an important element of flagellar resorption is destabilization of the microtubule-based axoneme, suggesting this signaling cascade may be evolutionarily conserved (Pan and Snell, 2005; Pan et al., 2004). Further supporting the idea of conservation, the C. elegans gene MEC-12 encodes an α-tubulin variant that is specifically required only in mechanosensing neurons, which depend on intact cilia: MEC-12 is the only α-tubulin in this species with a conserved site for acetylation (Fukushige et al., 1999). Interestingly, HDAC6 has been reported to associate with protein phosphatase 1 (PP1) (Brush et al., 2004), which binds microtubules (Liao et al., 1998), and dephosphorylates and inactivates AurA kinase. Such feedback may limit AurA activation at cilia.

A number of growth stimuli induce HEF1 expression and phosphorylation, influencing its protein interactions. These include PDGF, which is here shown to partially induce ciliary disassembly (Natarajan et al., 2006). Intriguingly, recent studies of p130Cas, a protein structurally similar to HEF1, indicate that p130Cas acts as a stretch sensor; HEF1 contains all sequence motifs necessary for similar function (Kostic and Sheetz, 2006). As one major function of cilium is to sense fluid flow, and overly persistent flow has been reported to induce ciliary disassembly (Iomini et al., 2004), stretch sensation may be an important action of HEF1. Our data suggest that HEF1 both activates AurA and stabilizes the protein from degradation; it will be interesting to determine if the HEF1 scaffolding activity also contributes to AurA interaction with its effector HDAC6. Our data also indicate that AurA activity influences IFT88 localization during disassembly, and suggest integrity of the IFT system is important for the disassembly process in animals, as in Chlamydomonas (Pan and Snell, 2005).

Our establishment of a HEF1-AurA-HDAC6 cascade at cilia also informs the understanding of the mitotic activities of these proteins. Dynamic changes in microtubule acetylation and deacetylation characterize the stages of mitosis, and HDAC inhibitors that inhibit family members with microtubule deacetylase activity induce mitotic arrest (Blagosklonny et al., 2002). The identification here of HDAC6 as an AurA target suggests that HEF1-AurA regulation of tubulin deacetylation at mitosis through HDAC6 might offer a mechanism to fine-tune the mechanical properties of the mitotic spindle. This signaling cascade may also influence re-establishment of focal adhesions at and following cytokinesis, given the growing appreciation of the role of microtubules in guiding the formation of these structures (Ezratty et al., 2005; Strickland et al., 2005). Further, one intriguing possibility is that the common use of an AurA-HEF1-HDAC6 switch at the basal body of quiescent cells and the centrosome of G2/M cells may serve as part of a checkpoint mechanism coordinating responsiveness to extracellular cues at different points in cell cycle. In this context, our observation that inhibition of AurA causes appearance of mitotically arrested cells possessing both spindles and cilia (results not shown) may reflect triggering of such a centrosomally based checkpoint.

These results also have implications for the understanding and treatment of cancer. Tumor cells commonly do not have cilia, and both HEF1 and AurA are often upregulated in cancer. The roles for these proteins at the centrosome and focal adhesions described earlier already offer two mechanisms by which these proteins may promote tumor initiation and progression. The current study indicates a third mechanism, in which elevation of HEF1 or AurA in tumors may destabilize cilia, thus conditioning cellular response to external cues and impacting multiple signaling pathways. Further, AurA is regarded as a promising chemotherapeutic target, with agents inhibiting this protein currently in clinical trials (Andrews, 2005). TSA and other broad-spectrum agents targeting HDACs are used in the clinic (Vanhaecke et al., 2004), with more focused agents such as tubacin in preclinical development (Hideshima et al., 2005). Our data suggest that AurA- or HDAC-targeted drugs may have previously unappreciated in vivo effects involving cilia, that may contribute to the observed efficacy and/or side effects of these agents.

PKD is one of the best-described cilia-related diseases (Wilson, 2001), with mutation of the cilia-localized polycystin proteins 1 and 2 (PKD1 and PKD2) responsible for the significant majority of PKD patients. p130Cas interacts directly with complexes containing PKD1 and PKD2, and also with nephrocystins, cilia-associated proteins that are mutated in a second renal cystic syndrome, nephronophthisis (Benzing et al., 2001). Although an association of HEF1 with these proteins has never been assessed, HEF1 is abundant in the kidney and conserves many protein interaction sequences with p130Cas. It is also tantalizing to consider that closer connections exist between dysplastic disorders leading to cysts and cancer than have previously been appreciated. One of the surprising results of a recent large study to analyze the cancer genome was the identification of the PKHD1 protein, a ciliary protein which is mutant in autosomal recessive PKD, as commonly mutated in colorectal cancer (Sjoblom et al., 2006). Overall, deregulated AurA/HEF1/HDAC6 signaling appears to have broad implications for studies of human development and disease.

Example 2 Generation of a Mouse Model to Study PKD

At present, there are numerous competing models to explain the basis for cyst formation, and the differences between the various syndromes associated with kidney cysts. Studies of the signaling changes that occur in PKD have identified anomalous function of pathways that affect proliferation, cell cycle, and apoptosis (Edelstein, C. L. (2005) Cell Cycle 4:1550-4). Downstream elements of these signaling pathways include the tumor suppressors PTEN, TSC2, and p53, the oncogenes Bcl-2 and Akt, and other important growth regulators such as mTOR (Shillingford, J. M. et al. (2006) PNAS 103: 5466-71). Current therapeutic strategies are attempting to exploit this information by using drugs that target the relevant processes and pathways, such as the use of caspase inhibitors to reduce apoptosis, and mTOR inhibitors to block cell proliferation (Tao Y., et al. (2005) PNAS 102: 6954-9). In some cases, these approaches are alleviating symptoms and slowing cyst growth. However, no highly effective disease management strategy currently exists.

In the past several years, new insights into cyst pathogenesis have come from the consideration of the possible role of defects in renal cilia (Benzing et al., 2006; Snell, W. J., et al. Cell (2004) 117:693-7)). This “ciliary hypothesis” is based on the recognition that the protein products of genes mutated in PKD (and other pleiotropic syndromes involving cyst formation) both localize to cilia and impact ciliary function. Understanding the regulation of cilia in PKD, should provide the basis for novel therapeutic approaches to PKD.

In the past two years, increasing attention has focused on the identification of other structural and signaling proteins associated with the cilium, the basal body, or the adjacent plasma membrane. Importantly, many proteins that have been identified as the genetic cause of human developmental defects associated with polycystic kidney disease, including polycystins 1 and 2 (encoded by PKD1, PKD2), fibrocystin (PKHD1), nephrocystins (NPHP1,3-5), and inversin (NPHP2), in each case localize to cilia. Additional disease-associated proteins localizing to cilia or basal bodies include the Bardet-Biedl Syndrome (BBS) proteins. Defects in BBS genes lead to kidney failure associated with renal cysts, and also loss of eyesight, obesity, and diabetes. Kartagener syndrome, characterized by reversed left-right symmetry (“situs inversus”) of the heart, stomach and liver, as well as additional defects, arises from ciliary dyskinesia (Carlen B., et al., (2005) Ultrastruct. Pathol. (2005) 29:217-20). Finally, targeted or spontaneous mutation in mice of other cilia-associated proteins including the kinesin motor KIF3A or of Tg737/polaris, both involved in IFT, results in similar syndromes (Siroky, B. J., et al., (2006) Am J Physiol Renal Physiol.; Cano, D. A., et al., (2004) Development 131:3457-67; Yoder, B. K., et al., (2002) Am J Physiol Renal Physiol. 282:F541-52; Nishimura, T., et al., (2004) Nat Cell Biol. 6:328-34).

These and other studies make it clear that defects in proteins affecting ciliary functions are a major cause of renal cysts in general, and PKD in particular, as well as other serious diseases.

Mechanistically, the role of cilia in development and in disease is not yet well-defined, although the field is advancing rapidly (FIG. 14). Cilia protrude from the apical cells into adjacent lumenal space. In this space, extracellular flow of liquid bends the cilia in the direction away from the flow. This induces a mechanosensation signaling response, in which opening of a transient receptor potential (TRP) membrane-associated cation channel releases a pulse of Ca²⁺ into cells. Polycystin 1 regulates the polycystin 2 TRP channel, and defects in these proteins (as well as other cilia-associated proteins) can cause defects in Ca²⁺ uptake. This Ca²⁺ signal propagates among neighboring renal cells within a tubule through gap junctions connecting the cells. Defects in other proteins such as Tg737/polaris cause additional defects involving IFT and signaling in cilia, in which the cilia are shortened; in mice defective for Tg737 (Oak Ridge Polycystic Kidney (orpk) mice), Ca²⁺ increase is clearly abnormal (Siroky et al. (2006); Nishimura et al., (2004). Extending ciliary function, cilia are present and important for the growth of some cells not thought to be regulated by flow: in these cases, the cilia are thought to act as chemosensors, with signal transducing proteins accumulating at the basal body.

While many of the downstream signaling components activated by Ca²⁺ are not yet well defined, the functional consequences of cilial bending for embryonic development are clearly profound. In development, cilial bending by extracellular fluid flow sends a polarity cue that conditions the future direction of cellular propagation. Inability to sense such flow is likely to underly the situs inversus observed in individuals mutant for some ciliary proteins such as inversin. In cell migration, fluid flow over cilia has been shown in some cases to contribute a polarity cue providing a direction for cell migration in development (Sawamoto K., et al. (2006) Science 311:629-32; Ciruna B., et al., (2006) Nature 439: 220-4). Within kidneys, flow sensing is thought to regulate proliferative response, such that defective sensing may cause overproliferation and cyst development. Separately, recently described connections between BBS proteins and the planar cell polarity (PCP) machinery imply that altered polarity of the cell division plane may cause dysplastic growth during maintenance of renal tubules, again leading to cyst formation (Fischer, E., et al., (2006) Nat. Genet. 38:21-3; Ross, A. J., et al., (2005) Nat. Genet. 37:1135-40).

An important point to consider is that as quiescent cells are induced to cycle, the cilium is reabsorbed, and the basal body returns to function within the centrosome, which includes action as microtubule organizing center (MTOC) for the bipolar spindle in mitosis. Although there are some differences among the cell systems used to study cell cycle regulation of cilia, most studies agree that cilia are reabsorbed by the time a cell enters mitosis, then re-form at several hours after the completion of cytokinesis (Quarmby et al., (2005). There are several implications of these findings. As discussed below, in addition to BBS, a number of other proteins have been identified at both basal bodies and centrosomes, and may act at both structures. These proteins may act directly at cilia and/or at centrosomes to orient the mitotic division plane of renal cells, allowing normal formation and maintenance of renal tubules. Conversely, as we demonstrate herein proteins that function at the centrosome to govern entry and exit from mitosis are ideally positioned to influence the formation and disassembly of cilia, and hence altered regulation of these proteins may contribute to the pathogenesis of PKD and related cilia-based syndromes.

Accordingly, the in vivo roles of AurA and HEF1 in cyst formation will be examined using mouse models. Appropriate models are available for this purpose. Notably, in late 2005, Seo et al. first described a HEF1 knockout mouse (Seo et al, (2005) J. Immunol. 175:3492-501). This knockout is viable and able to reproduce as a homozygote. To date, based on the interests of Sachiko Seo and her colleagues, characterization of these mice has been limited to the hematopoietic system. Even from this very limited analysis, it is clear that elimination of HEF1 has at least some phenotypic consequences, as B-cell maturation is defective. These mice will be utilized to explore the role of HEF1 in cell cycle and centrosome functions, and we have recently established a colony. Models for study of overexpressed AurA also exist (Fukuda et al., (2005) Mol. Cell Biol. 25:5270-81). However, in vivo overexpression of AurA has been reported as technically challenging, in part because of efficient proteasomal degradation of the protein. Further, AurA overexpression causes secondary phenotypes related to the failed cytokinesis seen in cells with too much AurA (Warner et al., (2003) Mol. Cancer Ther. 2:589-95). Instead, we will use small molecule inhibitors of AurA and HDACs to perform in vivo manipulation of these proteins.

Numerous mouse models have been developed for the study of PKD. Particularly because of the reported interaction of the HEF1-related protein p130Cas with polycystin, we wished to first explore the consequences of HEF1 and AurA in modulation of PKD1-associated PKD. A particularly attractive model is the conditional floxed Pkd1 model developed by the Germino group (Piontek K. B., et al. (2004) J. Am. Soc. Nephol. 15:3035-43). Mating of these animals with mice expressing Cre recombinase causes somatic loss of Pkd1, and leads to formation of renal and hepatic cysts by 10 weeks of age.

Taken as a whole, our results clearly indicate that HEF1 and AurA are regulated in time and space in a manner compatible with a controlling role in ciliary disassembly. They also demonstrate that increased activity and/or expression of HEF1 and AurA actively promote ciliary disassembly, and that clinical agents that block AurA stabilize cilia. Importantly, these results validate genetic predictions from Chlamydomonas CALK, demonstrating evolutionary conservation of AurA regulation of cilia and flagella. This conservation allows us to exploit ongoing discoveries regarding CALK in Chlamydomonas to guide our future studies. In the context of these and other published works, we theorize that a basal body-associated complex including HEF1 and polycystin (and potentially other proteins) comprises a stretch- and growth factor-responsive sensor at the cilium. In normal cells, AurA localizes to the basal body, but is only activated following receipt of signals through the HEF1-containing stretch complex. We will analyze the factors contributing to AurA activation. We also hypothesize that upon activation, AurA phosphorylates substrates located in the basal body and cilia. We also expect to observe moderately elevated expression of wild type or even inactive AurA influences the activity of effectors regulating ciliary disassembly, as well as cilia-associated signaling proteins. Methods are also provided to elucidate the mechanisms by which AurA and HEF1 condition cilia-associated signaling responses, and promote ciliary disassembly. Finally, as mentioned above, the consequences of modulating HEF1, AurA, and HDAC6 upon cyst formation in a PKD mouse model system will also be assessed.

To generate the required mouse strains for the experiments, we will first take the HEF1^(−/−) mice we have received from Seo and coworkers and backcrossed to a C57/B16 background. Our first step will be to create a series of mice that are heterozygous or homozygous null for HEF1 on a homozygous Pkd1^(cond/cond) background. See Piontek et al. In parallel, we will cross the HEF1 null mice to the nestin^(Cre) strain, ultimately creating mice heterozygous or homozygous null for HEF1, and hemizygously bearing the nestin^(Cre) gene. Next, we will combine these strains of mice to allow us to examine the consequences of HEF1 status on cyst formation in Pkd1^(cond/cond)nestin^(Cre) mice. These same mice can also be used for evaluation of the AurA and HDAC inhibitors. We emphasize, we do not view null status for HEF1 as independently likely to generate kidney cysts; rather, we expect the most likely activity will be to modify cyst formation induced by somatic mutation of Pkd1. See FIG. 15.

The central component of these experiments will be to compare rate and degree of cyst formation in HEF1^(−/−)Pkd1^(cond/cond)nestin^(Cre), HEF1^(wt/wt)Pkd1^(cond/cond)nestin^(Cre), and HEF1^(−/−)Pkd1^(wt/wt)nestin^(Cre) mice. We anticipate that most HEF1^(wt/wt)Pkd1^(cond/cond)nestin^(Cre) will have extensive cyst formation at 8-12 weeks of age. For each of the three strains of mice, we will sacrifice 10 animals at 8 and 16 weeks of age, and will use standard approaches to analyze timing, number, size, and pathological features of cyst formation. This will allow us to determine whether loss of HEF1 independently promotes cyst formation in the kidney, and whether lack of HEF1 positively or negatively regulates cyst formation dependent on defects in Pkd. We note, our crosses will also generate HEF1^(−/wt)Pkd1^(cond/cond)nestin^(Cre) animals: if a significant effect is seen with HEF1 null status, we will then determine whether HEF1 heterozygous status has an intermediate phenotype.

We note that there is (at present) no evidence directly implicating mutation of HEF1, its family member p130Cas, or Aurora in hereditary PKD in humans. This may still emerge. However, we view it as more likely that mutation of these genes would have a broader effect: for instance, mice with loss of p130Cas are embryonic lethal. By contrast, elevated levels of p130Cas, AurA, and HEF1 (unpublished) are found in numerous cancers, and thought to promote deregulated cell growth. We believe similar conditioning of cell growth by some of these proteins occurs in PKD. The methods disclosed herein will lead to the creation of new therapeutic strategies to treat PKD, by providing the scientific basis to apply existing compounds already in use for treatment of cancer to this disease. Moreover, the mice described herein will provide an in vivo model to assess the efficacy of various agents that may be useful for the treatment of diseases associated with aberrant cilia formation, such as PKD In addition, by improving our understanding of the molecular basis of PKD, we may be able to better predict disease progression and severity. Overall, the goal is to reduce the incidence and/or severity of PKD in those genetically prone to the disease, alleviate the symptoms of PKD in early stage patients, and limit the number of PKD patients that progress to end stage renal failure.

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Example 3 Rapid Transient Activation of Aurora-A Kinase Contributes to Interphase Calcium Signaling Responses

The Aurora-A (AurA) kinase is a member of the evolutionarily conserved Ipl family of kinases (reviewed in (Marumoto et al., 2005)). AurA is most abundant at the centrosome in G2 to M phase (Fukuda et al., 2005), and in studies performed in mammals and model organisms including Drosophila, AurA has been shown to be activated at mitotic entry, and perform critical functions in regulating entry into and passage through mitosis (Marumoto et al., 2005; Pugacheva and Golemis, 2006). In the past several years, AurA has attracted increasing attention because it has been found to be overexpressed in a high percentage of tumors arising in breast, colon, ovary, and other tissues (Bischoff et al., 1998; Goepfert et al., 2002; Tanaka et al., 1999; Tanner et al., 2000; Zhou et al., 1998), and because it has been shown to function as an oncogene when exogenously expressed in various cell line models (Anand et al., 2003; Meraldi et al., 2002; Tatsuka et al., 1998; Zhang et al., 2004). AurA is now being actively exploited as a target for development of new anti-cancer agents (reviewed in (Andrews, 2005)), based on its known role as a mitotic regulator.

Intriguingly, a number of studies have emerged in recent years to challenge the idea that AurA is solely a mitotic kinase. Serum induces AurA activation at the basal body of the cell cilium in non-cycling G0/G1 phase mammalian cells, causing AurA-dependent ciliary resorption (Pugacheva et al., 2007), and hence indirectly impacting the functionality of the cilia-dependent and cancer-relevant Hedgehog signaling cascades (Wong et al., 2009). AurA directly phosphorylates and regulates the activity of the RalA GTPase, an important EGFR/Ras effector important in many cancers (Wu et al., 2005), with this activity observed in interphase cells. AurA also has been reported to regulate microtubule dynamics in interphase cells (Lorenzo et al., 2009). All of these studies strongly imply a non-mitotic activity for AurA.

In a series of studies, we here demonstrate that interactions between the NEDD9 (also known as HEF1 and CAS-L) scaffolding protein and AurA are important for AurA activation at mitotic entry (Example I) and prior to ciliary resorption in interphase cells (Pugacheva et al., 2007). Interestingly, based on mRNA and protein analysis, NEDD9 (Law et al., 1996; Law et al., 1998) and AurA (Kurahashi et al., 2007) are predicted to be abundant in kidney. Pathologic conditions of the kidney include renal cell carcinoma (RCC), which has been linked to elevated AurA expression (Kurahashi et al., 2007). Abnormal cell division associated with formation of renal cysts is also very strongly linked to the changes in Ca²⁺ signaling induced by autosomal dominant polycystic kidney disease (ADPKD)-associated mutations in the PKD1 and PKD2 genes, encoding the PC1 transmembrane flow receptor and the PC2 calcium channel (Benzing and Walz, 2006; Pan et al., 2005; Wilson, 2004). Interestingly, an antibody cross-reactive with NEDD9 and its paralog BCAR1/p130Cas detected one of these proteins in a complex with PC2 (Geng et al., 2000). NEDD9 is also known to bind directly to the differentiation regulatory protein Id2 (Law et al., 1999), which in turn has been reported to bind directly to PC2 and mediate proliferative signals in PKD (Li et al., 2005). Further, recent studies of a PC2 ortholog in Chlamydomonas suggested a role for this protein in increasing intraflagellar calcium concentrations during the mating response (Huang et al., 2007), shortly prior to the activation of the CALK kinase. Separately, a recent study of Xenopus oocyte maturation indicated that inhibition of Ca²⁺ signaling led to eventual failure to accumulate and activate AurA (Sun et al., 2008). Cumulatively, these studies led us to hypothesize that NEDD9 and AurA might physically or functionally interact with PC2 to regulate cellular calcium signaling under normal or pathological conditions.

Based on these and other studies, we have investigated the role of AurA in calcium signaling responses. Our data here demonstrate that elevated cytoplasmic calcium signals rapidly and transiently through calmodulin to activate AurA, and that active AurA directly binds and phosphorylates PC2 to limit its Ca²⁺ channel activity, providing the first clear mechanism for AurA activation in non-mitotic cells. Our work also demonstrates AurA is abundant and frequently active in normal renal tissue, and hyperactivated in renal cysts associated with PKD. Finally, ADPKD affects as many as 1 in 500 individuals, and currently has few viable treatment options. We show that low concentrations of drugs that inhibit AurA activity augment PC2-dependent Ca²⁺ release, suggesting potential clinical applications in renal proliferation-associated pathologies.

The following materials and methods are provided to facilitate the practice of example 3.

Plasmids and constructs. Lentiviral constructs were obtained by cloning full-length PKD2 into the pLV-CMV-H4-puro-vector (provided by P. Chumakov and A. Ivanov). PKD2 cloned in pcDNA3.1-myc was provided by Dr. Stefan Somlo. The PKD2 CT fragment (amino acids 779-968) was cloned into the pEGFP (Clontech, Mountain View, Calif.) and pGEX-6P1 vectors (Millipore, Billerica, Mass.). Amino acid substitution mutations were introduced into wild type human PKD2 cDNA by site-direct mutagenesis, using the QuikChange XL Site-Directed Mutagenesis Kit (Stratagene, La Jolla, Calif.). Primer sequences are available on request. The FLAG-fused C-terminal domain of PKD1 (aa 4191-4302), containing the PC1-PC2 interaction site, was cloned in pcDNA3.1(+) vector (Invitrogen, Carlsbad, Calif.). FLAG- and GST-fused HEF1 were expressed from the vectors pCatch-FLAG (O'Neill and Golemis, 2001) and pGEX-2T (Law et al., 1998), respectively. AurA and derivatives were expressed from pCMV-SPORT6-C6 (OpenBiosystems, Huntsville, Ala.) and pcDNA3.1-mRFP vectors. A PCR product of mRFP1 was ligated into pcDNA3.1(+) (Invitrogen, Carlsbad, Calif.) to create pcDNA3.1-mRFP. pLV-CMV-H4-puro-vector, pEFGP and pcDNA3 were used for negative controls. Cell culture and transfection. HEK293 cells were maintained in DMEM with 10% FBS, plus penicillin/streptomycin. The immortalized human kidney proximal tubular cell line (HK-2, ATCC, catalog #CRL-219) was grown to subconfluence in keratinocyte serum-free media (Invitrogen, Carlsbad, Calif.). We transiently infected HEK293 cells with expression constructs for PKD2, HEF1 and AurA using Lipofectamine and Plus reagent (Invitrogen, Carlsbad, Calif.), according to the manufacturer's instructions. Cells were used for electrophysiological studies 24-48 hours post-transfection. For lentiviral infection, pLV constructs were co-transfected with pVSV-G and psPAX2 into the packaging cell line 293-T. After 24 h, media was collected, filtered through a 0.45-μm PVDF filter (Millipore, Billerica, Mass.), and applied to HK-2 cells with 8 μg/μl polybrene (Sigma, St Louis, Mo.) for 2 days, with fresh viral supernatant added every 12 h. After 48 h, cells were lysed, analyzed by western blot analysis, and used for further experiments. HK-2 cells stably expressing PC2 were obtained by infecting the HK-2 cell line with the pLV-PKD2 lentiviral vector, then selecting for 6-10 days with 1 mg/ml puromycin to produce a mass culture, as in (Pugacheva and Golemis, 2005). PC2 expression was verified by immunoblot and immunofluorescence analyses. Transient transfection of siRNAs was carried out using RNAi max transfection reagent (Invitrogen, Carlsbad, Calif.). Cells were assayed after 48 h of transfection. IC₅₀ determinations with the AurA kinase inhibitor PHA-680632 (Nerviano Medical Sciences, Nerviano, IT) were performed as described in (Skobeleva et al., 2007). Immunofluorescence. Cells growing on coverslips were fixed with 4% paraformaldehyde (10 min) then methanol (5 min), permeabilized with 1% Triton-X100 in PBS, blocked in 1×PBS with 3% BSA, and incubated with antibodies using standard protocols. Primary antibodies included mouse anti-AurA (BD Biosciences, San Jose, Calif.) and rabbit polyclonal anti-phospho-AuroraA/T288, (Cell Signaling, Beverly, Mass.), polyclonal anti-α-tubulin (Sigma), anti-α-tubulin mAb (Sigma), anti-acetylated α-tubulin mAb (clone 6-11B-1, Sigma, St Louis, Mo. and clone K(Ac)40 Biomol), anti-PC2 (G20, Santa Cruz Biotechnology, Santa Cruz, Calif., and YCC2 (a gift of S. Somlo), and mouse anti-protein disulfide isomerase (PDI) mAb (Abcam, Cambridge, Mass.). Secondary antibodies labeled with Alexa-488, Alexa-568, and DAPI to stain DNA, were from Molecular Probes/Invitrogen, Carlsbad, Calif. Confocal microscopy was performed using a Nikon C1 Spectral confocal microscope (Nikon). Protein expression, western blotting and immunoprecipitation. Recombinant glutathione-S-transferase (GST), GST-fused to amino acids 779-968 of the PC2 carboxy-terminus (GST-PC2₇₇₉₋₉₆₈) and NEDD9 (aa 1-363, previously shown to bind and activate AurA, (Pugacheva and Golemis, 2005)) were expressed in BL21 (DE3) bacteria, induced with IPTG, and purified using the MicroSpin GST Purification module (GE Healthcare, Piscataway, N.J.). Purified recombinant AurA was purchased from Upstate (Charlottesville, Va.). For western blotting and immunoprecipitation, mammalian cells were disrupted in CelLytic M lysis buffer (Sigma, St. Louis, Mo.) supplemented with a protease and phosphates inhibitor cocktails (Roche, Basel Switzerland). Whole-cell lysates were used either directly for SDS polyacrylamide gel electrophoresis (SDS PAGE), or for immunoprecipitation. Immunoprecipitation samples were incubated overnight with antibody at 4° C., and subsequently incubated for 2 h with protein A/G-sepharose (Pierce, Rockford, Ill.), washed and resolved by SDS-PAGE. For analysis of PC2 glycosylation, cell lysates were treated with endoglycosidase H (Endo H) (New England Biolabs, Beverly, Mass., USA) and analyzed by SDS-PAGE followed by immunoblotting as described previously (Cai et al., 1999; Koulen et al., 2002). GST-pulldown assays used wild type AurA (Upstate, Charlottesville, Va.) mixed with titrated quantities of GST and GST-PC2₇₇₉₋₉₆₈. Western blotting was done using standard procedures, and developed by chemoluminescence using the West-Pico system (Pierce, Rockford, Ill.). Primary antibodies included mouse anti-NEDD9 mAb (clone 2G9, (Pugacheva and Golemis, 2005)), anti-AurA (BD Biosciences, San Jose, Calif.), anti-phospho-AurA-T288 (Cell Signaling, Beverly, Mass.), anti-Myc- and anti-VlaR (Santa Cruz Biotechnology, Santa Cruz, Calif.), anti-β-actin mAb (AC15, Sigma, St Louis, Mo.), and anti-PC2 (G20, Santa Cruz Biotechnology, Santa Cruz, Calif., and YCC2, a gift of S. Somlo). Rabbit anti-GFP (Abcam, Cambridge, Mass.; ab290) was used for immunoprecipitation, and mouse anti-GFP (JL-8; BD Biosciences, San Jose, Calif.) was used for western blotting. Anti-GST mAb (Cell Signaling, Beverly, Mass.), polyclonal EZview Red anti-Flag M2 affinity gel (Sigma, St Louis, Mo.) and polyclonal anti-AurA agarose immobilized conjugate (Bethyl, Montgomery, Tex.) were used for immunoprecipitations. Secondary anti-mouse and anti-rabbit HPR conjugated antibodies (GE Healthcare, Piscataway, N.J.) were used at a dilution of 1:10,000 for visualization of western blots. Image analysis was done using NIH ImageJ—Image Processing and Analysis software (National Institutes of Health, Bethesda, Md.), with signal intensity normalized to β-actin or total AurA level.

To assess AurA phosphorylation of PC2 in vivo, myc-tagged PC2 was transiently expressed alone or with AurA or T288D-AurA in HEK293 cells and then immunoprecipitated with anti-myc antibody (Santa Cruz Biotechnology, Santa Cruz, Calif.). Phosphorylation of the S829 site was assessed by western blot with Phospho-(Ser/Thr) PKA Substrate Antibody (Cell Signaling, Beverly, Mass.). 500 nM PHA-680632 (Nerviano Medical Sciences, Nerviano, IT) was used to inhibit AurA and 10 μM H89 PKA inhibitor (Calbiochem) were used to inhibit phosphorylation.

Calmodulin pull-down assay and Immunoprecipitation. Cell lysates in lysis buffer (PBS with 1% Triton X-100) or purified GST fusion protein diluted in binding buffer (50 mM Tris-HCl, pH7.6; 120 mM NaCl; 1% Brij) were incubated with Calmodulin-Sepharose® 4B (GE Healthcare, Piscataway, N.J.) or control Sepharose for 1-2 h at 4° C. as indicated in the figure legends. After washing, beads were boiled in SDS sample buffer and separated by SDS-PAGE followed by Western blotting. Kinase assays. To assess phosphorylation of PC2 by AurA, an in vitro kinase assay was performed using bacterially expressed GST-fused PC2 CT and recombinant active AurA (Upstate, Charlottesville, Va.) or overexpressed AurA immunoprecipitated from mammalian cells. in standard kinase buffer with addition of an Mg/ATP cocktail (Upstate, Charlottesville, Va.). MBP (Upstate, Charlottesville, Va.) and histone H1 (Upstate, Charlottesville, Va.) were used as positive and negative controls for AurA phosphorylation, using standard methods. Parallel aliquots without γ-³²P(ATP) were processed for SDS-PAGE/Coomassie staining (Invitrogen, Carlsbad, Calif.). To assess CaM-dependent AurA activation in vitro kinase assay was performed using AurA purified from baculovirus or according the protocol described above in the presence of 1 μM CaM (Calbiochem) and 1 mM Ca²⁺. For kinase assay without Ca²⁺, 1 mM EGTA was substituted for 1 mM Ca²⁺ in the kinase buffer. Mass spectrometry. After an in vitro kinase reaction with AurA produced from baculovirus in the presence or absence of CaM plus 1 mM CaCl₂, gels were stained with Coomassie blue, and phosphorylated AurA bands were excised and sequenced at the Taplin Biological Mass Spectrometry Facility at Harvard Medical School, by using microcapillary reverse-phase HPLC nanoelectrospray tandem mass spectrometry on a Finnigan LCQ DECA quadrupole ion trap mass spectrometer). Cytosolic Ca²⁺ measurements. Cells expressing PC2 constructs were plated on glass coverslips and grown to ˜80% subconfluence. The coverslips were rinsed in Hanks' balanced salt solution (HBSS), and incubated with Fluo-4AM (5 μM in HBSS) (Invitrogen, Carlsbad, Calif.) in the presence of 0.02% pluronic acid (Invitrogen, Carlsbad, Calif.), and 2.5 mM probenecid (Invitrogen, Carlsbad, Calif.) for 15-30 minutes at RT. The coverslips were washed twice in HBSS, mounted in a perfusion chamber (FC2, Bioptechs, Butler, Pa.), and analyzed with a Nikon C1 Spectral confocal microscope (Nikon). Cytosolic Ca²⁺ measurements done in the absence of extracellular Ca²⁺ were performed on cells washed and assayed in the HBSS described above except that CaCl₂ was omitted and 0.5 M EGTA was added. In experiments involving AurA inhibition, cells were treated for 2 h with 500 nM PHA-680632 (Nerviano Medical Sciences, Nerviano, IT) before calcium measurement. Fluo-4 was excited at 488 nm and emission was time-lapse recorded at 522 nm. Cells were individually selected, and their fluorescence intensities (F) were normalized to baseline (F0), and analyzed with Metamorph and Metafluor softwares (Molecular Devices, Union City, Calif.).

To compare cellular responses, we examined differences in intensity over time using a generalized linear model assuming Gamma distribution and log link. We fit the models by generalized estimating equations assuming an autoregressive correlation structure to account for correlation of observations over time. We included baseline intensity, group, time, and the interaction between time and group in the models. To allow for flexible effects over time, we entered time and the related interactions in the model using restricted cubic splines assuming 5 knots (Harrell, 2001). We used Wald tests to assess p-values of group effects at each time point.

To induce ciliary formation in the HK2 cell line, cells were plated at 50%-70% confluence and grown for 3-4 days in medium with 10% serum, then grown for 2 days in Opti-MEM medium without serum. Immunofluorescence was used to confirm >80% of HK2 cells were ciliated before use in experiments. Immunohistochemistry. All tissue samples examined were IRB-consented, 10-20 mm sections of formalin-fixed, paraffin-embedded tissues representing either normal human kidney tissue, or kidney tissue from patients diagnosed with PKD. A standard two-stage indirect immunoperoxidase staining protocol was used for all tissues (Vectastain ABC System; Vector Laboratories), with antigen retrieval buffer from BD Biosciences, San Jose, Calif. As controls, sections were stained with diluent alone (5% goat serum in Tris-buffered saline) and antibody pre-absorbed with the immunizing peptide. Incubations with tissue sections were carried out at room temperature for 1 h or at 4° C. overnight, and subsequent steps were carried out at room temperature. Staining was visualized with either diaminobenzidine (brown) or VIP (purple) (Vector Laboratories), and sections were counterstained with hematoxylin (Sigma, St Louis, Mo.). Antibody to total AurA (anti BTAK ab, provided by T. Gritsko) was used at a dilution of 1:500, and to phospho-T288 AurA at 1:100 (Bethyl, Montgomery, Tex.). Images were acquired at 10× and 40× using a Nikon Eclipse E600 microscope. Statistical Analysis. Statistical comparisons were made using a two-tailed Student's t test. Experimental values were reported as the means±S.E. Differences in mean values were considered significant at p<0.05. All calculations of statistical significance were made using the GraphPad InStat software package (San Diego, Calif.).

Results

Rapid transient activation of AurA accompanies stimuli for Ca²⁺ release from ER stores in kidney cells. Polycystin-2 (PC2) Ca²⁺ channel activity is stimulated by treatment of kidney cells with the peptide hormone and anti-diuretic arginine vasopressin (AVP) (Cai et al., 2004; Koulen et al., 2002), which signals through the AVPR1 receptor to initiate InsP3 and other second messenger cascades that initiate Ca²⁺ release from the ER. To assess whether activation of AurA occurred in this physiological context, we treated the HK-2 proximal tubule-derived human kidney cell line (Ryan et al., 1994) with AVP, and used immunofluorescence to measure AurA activation (FIG. 16A). Strikingly, T288-autophosphorylated AurA at centrosomes increased 3-fold within 5 seconds of AVP treatment, returning to basal levels after ˜2 minutes. AurA activation was induced by release of Ca²⁺ from internal ER stores, as equivalent results were obtained in cells cultured in medium with (results not shown) and without (FIG. 16A) Ca²⁺. As a complementary approach to confirm that PC2 activators induce rapid AurA autophosphorylation, we treated AurA-transfected HEK293 cells grown in Ca²⁺-free medium with either AVP or histamine, an InsP3 generating agonist that triggers activation of ER-associated PC2 (Brini et al., 2005; Koulen et al., 2002). In each case (FIG. 1B), T288-AurA appeared rapidly after stimulation, in cells grown in Ca²⁺-free medium.

The rapid AurA activation following treatment might be induced by the second messengers transmitting signals from membrane localized receptors, or alternatively, the elevated cytoplasmic Ca²⁺ levels resulting from ER-release might more directly regulate AurA activation, with no second messengers required. To distinguish between these mechanisms, we treated AurA-transfected HEK293 cells with the Ca²⁺ uptake inhibitor thapsigargin, which blocks the recycling of cytoplasmic Ca²⁺ into ER stores (Sagara and Inesi, 1991), thus increasing cytoplasmic Ca²⁺ levels. Thapsigargin treatment caused a rapid (<1 minute) increase in levels of activated T288-AurA, and increased the in vitro kinase activity of immunoprecipitated AurA against defined substrates (FIGS. 16C, 16D). The BAPTA-AM and EGTA Ca²⁺ ion chelators block Ca²⁺-mediated signaling pathway activation (Anyatonwu et al., 2007; Paria et al., 2006). BAPTA/AM used alone or at low concentration in combination with EGTA completely blocked thapsigargin-induced T288 AurA autophosphorylation (FIG. 16E). In contrast, EGTA alone, which chelates only extracellular Ca²⁺, had no effect, even in cells grown in medium containing Ca²⁺ (not shown). Moreover, treatment of AurA-transfected HEK293 cells grown in Ca²⁺-free medium with the Ca²⁺-selective ionophore ionomycin, which directly triggers Ca²⁺ release from the ER, similarly induced transient activation, with similar kinetics. Direct rapid activation of AurA by Ca²⁺ has not been previously demonstrated in mammalian cells, although the AurA partner protein and activator NEDD9 has been shown to be rapidly tyrosine-phosphorylated (associated with greater activating capacity for some partner proteins) after Ca²⁺ release in osteoclasts (Zhang et al., 1999; Zhang et al., 2002). However, in cells transfected with siRNA depleting NEDD9, thapsigargin induced comparable AurA phosphorylation to that in control transfected cells, suggesting that calcium activated AurA phosphorylation is independent from NEDD9 activity (FIG. 16F).

Calcium induces calmodulin to bind and activate AurA. Increased cytoplasmic Ca²⁺ might induce AurA activation by direct binding, or through inducing conformational changes in AurA by triggering its binding to a Ca²⁺-binding effector such as calmodulin (CaM), or by activating of an intermediate kinase or phosphatase that targets AurA, In an in vitro kinase assay, titration of Ca²⁺ into the reaction did not affect AurA autophosphorylation or activity towards substrates (not shown). However, addition of Ca²⁺ with CaM strongly induced AurA autophosphorylation (FIG. 17A), as well as AurA phosphorylation of NEDD9 and other substrates (FIGS. 17A, 17B). A small molecule inhibitor of AurA, PHA680632 (Soncini et al., 2006), blocked these CaM-induced phosphorylations (FIG. 17B). The direct AurA-CaM binding predicted by these results was confirmed both in vitro and in vivo (FIGS. 17C, 17D). Moreover, thapsigargin-induced AurA activation was completely blocked by the CaM inhibitor calmidazolium, both in HK2 and in HEK293 cells (FIG. 17E). Using mass spectrometry, we determined that addition of CaM to AurA resulted in the appearance of a substantial S51-phosphorylated AurA species (QRVLCP-S(51)-NSSQR). Interestingly, phosphorylation of the analogous S53 of the Xenopus AurA ortholog has previously been shown to be a product of mitotic auto-phosphorylation, and to stabilize AurA from proteasomal degradation at mitotic exit (Littlepage and Ruderman, 2002; Littlepage et al., 2002). Aurora A interacts directly with PC2. To assess the role of Ca-dependent AurA activation, and identify its physiological substrates, we considered a possible connection to PC2, based on the reported association between polycystins and Cas proteins (p130Cas and NEDD9). The bulk (>95%) of intracellular PC2 associates with the ER, and mediates Ca²⁺ release to the cytoplasm (Cai et al., 1999). To assess whether there might be direct interactions between AurA, its previously defined partner and activator NEDD9 (Pugacheva and Golemis, 2005), and PC2, we first established that endogenous AurA and PC2 coimmunoprecipitated from HK-2 cells (FIG. 18A). The PC2 C-terminus contains the primary sites for interaction with the mechanosensor PC1 (a.a. 832-895), as well as other important functional motifs. To assess AurA interactions with this PC2 domain, we next co-transfected AurA with the GFP-tagged PC2 cytoplasmic C-terminal domain (PC2-CT, amino acids 779-968) into HEK293 kidney cells. Overexpressed AurA and PC2-CT coimmunoprecipitated (FIG. 18B). Further, in a defined in vitro system, GST-fused PC2-CT pulled down recombinant purified full length AurA (FIG. 18C), interacting separately with both the AurA regulatory and catalytic domains (FIG. 18D). AurA did not compete with PC1 for binding to PC2 (FIG. 18E), suggesting the utilization of distinct binding sites on PC2. Finally, in contrast to the results with AurA, while overexpressed NEDD9 and PC2 coimmunoprecipitated (FIG. 18F), no interaction was seen between endogenous NEDD9 and PC2, or the two purified proteins in vitro system (not shown). Together, these data suggested a strong and direct interaction between AurA and the PC2 C-terminus, and a significantly weaker or indirect interaction between NEDD9 and PC2. AurA phosphorylates PC2 on C-terminal residue S829. We next asked if PC2 might be a substrate of AurA. Besides the PC1 interaction motif, the PC2 C-terminus (FIG. 19A) encompasses an EF hand, Ca²⁺ binding motif (a.a. 754-782), and endoplasmic reticulum (ER) targeting sequences (a.a. 787-820) (Giamarchi et al., 2006). A S812 casein kinase II (CKII) phosphorylation site is important for positively regulating PC2 Ca²⁺ channel activity (Cai et al., 2004). We identified a strongly consensus candidate AurA phosphorylation motif (Ferrari et al., 2005) at residue S829 (RRGSI), adjacent to the ER-targeting domain, and a less favorable motif at residue S944 (PRSSR). We established that recombinant, activated AurA phosphorylated the PC2 C-terminus in vitro (FIG. 19B). AurA phosphorylation of PC2 was enhanced by interactions with NEDD9 (Pugacheva and Golemis, 2005), in contrast to AurA phosphorylation of the control substrate MBP, which was unaffected (FIG. 19C). AurA phosphorylation of PC2 was separately enhanced by inclusion of CaM and Ca²⁺ in in vitro reactions (FIG. 19D). We next compared the ability of AurA to phosphorylate a wild type PC2 C-terminus, versus derivatives with S->A mutations in the S829, S944, or CK2 motifs, or combinations of these mutations (FIGS. 19E, 19F). An S829A mutation completely eliminated AurA phosphorylation of PC2, while S944A and S812A had no effect on this phosphorylation, either independently or in combination with S829A. By comparison, CK2 phosphorylated solely the S812 residue, and its phosphorylation of this site was not affected by the presence of S829A or S944A mutations (FIGS. 19G, 19H).

To investigate the in vivo phosphorylation of the S829 site, we exploited the fact that this site is quite similar to the general PKA substrate consensus (RRxS). Phospho-PKA substrate antibody recognized PC2 but not S829A-mutated PC2 in transiently transfected HEK293 cells (FIGS. 19I, 19J). Importantly, co-transfection of constitutively active AurA (T288D) increased phosphorylation of this site, while treatment of cells with an AurA inhibitor (PHA680632) but not a PKA inhibitor (H89) reduced in vivo phosphorylation. Those data indicate that full length PC2 is phosphorylated in vivo at S829.

AurA negatively regulates PC2 Ca²⁺ channel activity. We next asked if AurA might play a role in regulating the PC2 Ca²⁺ channel. We transiently transfected HEK293 cells with PC2 together with RFP-AurA or an RFP negative control, and measured fluorescence of the cytoplasmic Ca²⁺-binding dye Fluo-4 following AVP treatment, comparing the amplitude and duration of Ca2+ release (FIGS. 20A, 20B). Cotransfected RFP-AurA reduced the amplitude of Fluo-4 signal to 50% of that in control-transfected cells. An equivalent response was seen in cells in which histamine was used to induce PC2 channel activity. Reciprocally, treatment of PC2-transfected HEK293 cells with the AurA inhibitor PHA 680632 (Soncini et al., 2006) significantly enhanced the amplitude of release (FIGS. 20C, 20D). Similar results were obtained using HK2 cells stably overexpressing PC2 (FIGS. 20E, 20F), and in cells treated with a separate small molecule inhibitor of AurA, C1368. Finally, in PC2-overexpressing HK-2 cells with AurA depleted by siRNA, AVP induced Ca²⁺ release was significantly increased, to a degree comparable to that seen with treatment with AurA inhibitory drugs (FIGS. 20G, and 20H). Together, these data strongly implicated AurA as a regulator of PC2 channel activity. A cytoplasmic pool of AurA regulates ER-localized PC2. AurA is typically thought of as localized to the centrosome or centrosomally-derived ciliary basal body, which would limit access to an ER-localized pool of PC2. In contrast, cancerous cells with overexpressed AurA have an extensive diffuse pool of cytoplasmic AurA. However, AurA is intrinsically abundant in kidney cells, and indeed, siRNA depletion experiments followed by detection with two different antibodies to AurA (FIG. 21A and not shown) indicated the presence of a significant cytoplasmic pool of AurA in HK2 and other kidney cells, in addition to the anticipated concentrated pool of AurA at centrosomes and the ciliary basal body.

PC2 activity can be controlled by direct changes in the activity of the ER-localized protein, or alternatively, based on phosphorylation-mediated relocalization of PC2 between ER and other cellular compartments such as the cilium or plasma membrane, for example via CK2 phosphorylation at S812 (Kottgen et al., 2005). The experiments described above were performed in cycling, predominantly unciliated cells, leading us to favor the former mechanism; however, to specifically assess whether the presence of intact cilia influence AurA regulation of PC2 activity, we directly compared the ability of PHA-680632 to increase AVP-induced PC2 channel activity in predominantly non-ciliated (<15%), cycling versus ciliated (>80%), non-cycling HK-2 cells. These results showed no significant difference in the degree of Ca²⁺ release under the two growth conditions (FIG. 21B). Moreover, AurA phosphorylation did not inhibit PC2 localization to the ER. Cells cultured with or without the AurA inhibitor PHA-680632 showed comparable pools of endoglycosidase-H cleavable PC2, whether in ciliated or non-ciliated HK-2 cells (FIG. 21C), while PHA-680632 treatment did not increase the detection of PC2 at the cilia in these cells; however, we note that only approximately 10% of cilia in HK-2 cells stained positively for PC2 regardless of drug treatment, confirming this is a minor site of localization of the protein in this cell type. We next analyzed the consequences of mutating the S829 AurA phosphorylation site on PC2 expression, localization, and activity. Based on immunofluorescence (FIG. 21D) and endo-H cleavage (FIG. 21E), the degree of localization of both S829A and S829E derivatives to the ER is comparable to that of wild type PC2, supporting data obtained with the AurA inhibitor. Co-overexpression of AurA with PC2 also did not affect this ER localization (FIG. 21E). However, mutation of the AurA phosphorylation site significantly affected ER integrity, with cells expressing these mutants developed an abnormal, aggregated morphology within 24 hours (FIG. 21D), and dying within 48-72 hours of transfection or transduction.

Abundant AurA expression and activity in normal and cystic kidney tissue. If AurA has in vivo, non-cell cycle function relevant to PC2, AurA should be detectable and potentially active in non-dividing renal tissue. Immunohistochemical analysis of primary human kidney specimens readily detected AurA in multiple sub-structures (FIG. 22A). AurA was most concentrated in cells of the proximal and particularly distal convoluted tubules, and in the collecting ducts. AurA was not detectable in the glomerulus, or in the loops of Henle. This expression pattern is similar to that previously reported for PC2 (Foggensteiner et al., 2000). AurA staining was generally detectable in the cytoplasm, as seen in cultured kidney cells, but also intensely concentrated in the nucleus of some cells, with the greatest number of nuclear-staining cells associated with the distal convoluted tubules. Suggestively, a subset of these AurA-positive structures also stained positively for T288-phospho-AurA, indicating activity across adjacent groups of non-mitotic cells (FIG. 22A); again, the most intense staining was associated with distal convoluted tubules and collecting ducts. Further examination of 5 primary cysts derived from patients with PKD (FIG. 22B) revealed prominent AurA staining specifically within the epithelial cells lining the cyst, but not the fibrotic tissue, suggesting AurA expression is retained in renal cysts, and activity is increased over that in normal renal tissue. PHA-680632 enhances PC2 activity at low IC values. There is a considerable need for novel therapeutics to treat PKD. Given that one consequence of mutations in PKD1 is to reduce PC2 activation (Nauli et al., 2003), if AurA inhibition enhances PC2 activation, this might lead to therapeutic benefits. In vivo, Aurora kinase inhibitors have marked effects as cell cycle inhibitors (Gautschi et al., 2008), relevant to their action in cancer therapy, raising the possibility of toxic side effects if these agents used in PKD. However, some recent studies have suggested that the cytotoxic effects of Aurora kinase inhibitors used in vivo at least partially reflects their cross-reactive inhibition of Aurora-B rather than AurA, which occurs at higher concentrations (discussed in (Gautschi et al., 2008)).

We have compared the doses of PHA-680632 required to inhibit cell growth with those required to enhance PC2 signals. In HK-2 cells, the IC₅₀ value for PHA-680632 is 3.25 μM, while the 500 nM concentration used for the experiments described above represents an IC₅₀ value (FIG. 22C). In contrast, an approximately two-fold enhancement of AVP-induced Ca²⁺ release is seen whether PHA-680632 is used either at 3.25 μM or at 500 nM (FIG. 22D, 22E). One possible explanation for the difference in results could be that PHA-680632 was used for only 2 hours pre-treatment in Ca²⁺ release experiments, but must be sustained in culture medium for 3 days in IC₅₀ determinations: greater compound decay in the latter experiments might stipulate higher initial dosing concentrations. However, in parallel experiments in which PHA-680632 was added to media 2 or 24 hours before AVP treatment, significantly greater enhancement of PC2 activity occurred with the longer 24-hour pre-incubation than the 2-hour incubation used for other experiments, suggesting drug stability was not an issue (FIGS. 22D, 22E). These data suggest that AurA may be a useful target in modulating PC2 activity in vivo.

Discussion

The work presented here reveals and mechanistically defines a novel activity of AurA, as a direct target of calcium activation, and in direct control of cellular homeostasis for calcium. FIG. 23 represents a model for the data presented here. In this model, transient stimuli such as AVP or histamine trigger Ca²⁺ release to the cytoplasm, inducing calmodulin binding and auto-activation of AurA, marked by AurA S51 phosphorylation. Activated AurA phosphorylates ER-localized PC2 on 5829, restricting its Ca²⁺ channel activity, and then itself becomes inactive in the context of reduced cytoplasmic Ca²⁺. NEDD9 contributes to the specific interaction of AurA with PC2, likely by promoting the ability of AurA to phosphorylate PC2. Inhibition of AurA by small molecule inhibitors enhances PC2 activity, increasing the magnitude of ER Ca²⁺ release induced by upstream activators. Overexpression of AurA has the opposite effect, limiting PC2 channel activity.

Although to our knowledge a direct AurA-calcium connection has never previously been investigated, there are some suggestions that a connection between AurA and calcium signaling may be evolutionarily conserved. Flagellar disassembly in Chlamydomonas is induced by stimuli including gamete fusion following mating (Cavalier-Smith, 1974). Intriguingly, recent studies of a PC2 ortholog in Chlamydomonas suggested a role for this protein in increasing intraflagellar calcium concentrations during the mating response (Huang et al., 2007), prior to the activation of the CALK kinase. Separately, a recent study of Xenopus oocyte maturation indicated that inhibition of Ca²⁺ signaling led to eventual failure to accumulate and activate AurA, and subsequently a defective phenotype in meiosis (Sun et al., 2008). Dynamic changes in calcium signaling play a key role in meiosis, have been implicated in action at spindle-associated microdomains in mitosis (Parry et al., 2005), and may both regulate and be regulated by AurA. An attractive feature of Ca²⁺-dependent AurA regulation is that it offers a potential mechanism to help explain the rapid, timed activation of AurA at cell cycle transitions. At present, although multiple proteins have been demonstrated to bind and support AurA activation at the mitotic boundary, most of these also interact with AurA in G2, implying the existence of a triggering event at the actual transition point. The transient increase of cytoplasmic Ca²⁺ could provide a suitable mitotic trigger. Such Ca²⁺- and CaM-dependent regulation of the centrosomal proteins centrin and CP110 has been shown to be critical for the action of these proteins in supporting cytokinesis. In addition, reports that PC2 itself localizes to the mitotic spindle in dividing cells (Rundle et al., 2004) supports the idea that the mechanism of PC2-dependent Ca²⁺ release and calmodulin binding may be a physiological trigger for AurA during cell cycle, causing conformational changes that are permissive for other AurA interacting proteins to sustain its activity. However, the extremely transient nature of the Ca²⁺-dependent activation we observe makes it difficult to test this idea directly.

Interestingly, published studies of AurA in cancer often suggest the pro-oncogenic activity of AurA may arise because its overexpression allows a normally centrosomal protein access to inappropriate substrates. However, our results suggest changes in AurA expression in cancer may promote quantitative changes in AurA activity more than qualitative phosphorylation of novel substrates, as based on the data presented here, AurA has a broader sub-cellular localization profile in untransformed cells than is commonly appreciated, with the centrosome acting as a concentration point at which the protein is most readily visualized. Our data indicate that not only kidney cells, but also primary kidney tissue express significant quantities of AurA in the cytoplasm and nucleus, and that some of this AurA is activated in non-cycling cells in normal kidney tissue, particularly in cells of the distal convoluted tubules and collecting ducts from which cysts arise. This broader view of AurA activity is compatible with recent reports that AurA phosphorylates RalA (Wu et al., 2005) (a protein not known for either centrosomal localization or mitotic-specific action, but which has itself been described as calmodulin-binding (Clough et al., 2002)).

In general, calcium signaling differs significantly in cancerous (Roderick and Cook, 2008) and cystic (Harris and Torres, 2008) cells versus normal cells, promoting increased cell proliferation through the abnormal activation of numerous calcium-responsive signaling pathways. The fact that AurA activation was elevated in PKD-associated cysts is interesting, and may reflect paradoxical activation in the context of mutated PKD1 and PKD2, analogous to the overexpression of growth inhibitory proteins in tumors that have eliminated partners in a feedback loop. Based on our results, it appears that inappropriately activated AurA may act as an intermediate in some of signaling processes relevant to PKD. For example, besides binding the PC2 partner Id2, NEDD9 directly binds and is both target and activator of Src kinase (recently discussed in (Singh et al., 2007)). Src signaling is abnormal in PKD, and a recent study has indicated that inhibition of Src produces clinical benefits in PKD (Sweeney et al., 2008). Through interactions with NEDD9, AurA may influence the activity of Src and Id2 in either normal renal tissue or in cysts. These close physical interactions suggest further topics of study not only in renal cysts but also in cancer, where NEDD9, Src, and Id2 all have oncogenic function.

There is potential therapeutic benefit to identifying ways to stimulate PC2 channel activity, and there is an urgent need to develop effective therapies for PKD. At present, a number of targeted therapeutic agents are moving through pre-clinical development and clinical trials. Besides c-Src, these include agents targeting mTOR, HER2, and others. These studies provide a precedent for adapting drugs originally developed as cancer therapeutics in PKD. An obvious concern is that given the chronic but survivable nature of PKD, it is necessary to be extremely cautious in using powerful compounds that may themselves ultimately select for oncogenic changes. However, our data suggest that very low doses of an AurA-targeting inhibitor are able to enhance PC2 activity, suggesting a basis for further investigation of such agents in cases of PKD linked to PKD1 mutation, where PC2 is insufficiently activated but structurally intact. It is also fascinating to note that defects in PKD1 and PKD2 have recently been linked to centrosomal amplification in both animal models and human patients (Battini et al., 2008; Burtey et al., 2008), reducing the separation between cystic syndromes and cancer, and supporting the idea that calcium-dependent activation of AurA is relevant to the severity of PKD presentation.

It is clear that mutations that cause polycystic kidney disease (PKD) reduce the functionality of two proteins, PC1 and PC2, encoded by the PKD1 and PKD2 genes. Therapies that help increase PC1 and/or PC2 activity are predicted to help reverse the consequences of having PKD1 or PKD2 mutations. Our group has for the first time identified a role for the Aurora-A kinase in negatively regulating PC2 activity. Because Aurora-A has attracted interest as an oncogene important for some cancers, drugs have been developed and are being evaluated in clinical trials that can block Aurora-A activity. The data provided herein indicate that these drugs increase PC2 function, thus supporting their use as new therapeutics for PKD. See FIG. 23.

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While certain preferred embodiments of the present invention have been described and specifically exemplified above, it is not intended that the invention be limited to such embodiments. Various modifications may be made to the invention without departing from the scope and spirit thereof as set forth in the following claims. 

1. A method for identifying agents which modulate calmodulin-Aurora A complex formation having efficacy for the treatment of polycystic kidney disease (PKD), comprising: a) providing kidney cells which express Aurora A and calmodulin and incubating cells in the presence and absence of said agent and in the presence of a molecule which induces release of Ca²⁺; b) determining the extent said agent disrupts calmodulin-Aurora A complex formation following release of calcium, agents which disrupt complex formation having efficacy for the treatment of PKD.
 2. The method of claim 1, wherein said agent inhibits activation of aurora A kinase.
 3. The method of claim 2, wherein said aurora A kinase inhibitor is selected from the group consisting of siRNA which specifically down modulate Aurora A expression, C1368, PHA-680632, MK-0457, ZM447439, MLN 8237, MLN 8054, VX-680 and hesparadin.
 4. The method of claim 1, comprising determining whether said agent disrupts aurora A binding to PC2.
 5. The method of claim 1, comprising determining whether said agent alters cellular localization of PC2.
 6. The method of claim 1, comprising determining whether said agent alters Aurora A phosphorylation of PC2.
 7. A method for inhibiting progression of polycystic kidney disease in a patient in need thereof, comprising administration of an effective amount of an aurora kinase inhibitor, said inhibitor being effective to inhibit cyst formation in the kidney.
 8. The method of claim 7, wherein said kinase inhibitor is effective to inhbit aurora kinase A and aurora kinase B.
 9. The method of claim 7, wherein said kinase inhibitor is effective to inhibit aurora kinase A or aurora kinase B.
 10. The method of claim 7, wherein said aurora kinase inhibitor is selected from the group consisting of siRNA which specifically down modulate Aurora A expression, C1368, PHA-680632, MK-0457, ZM447439, MLN 8237, VX-680 and hesparadin. 